Process of making a compound by forming a polymer from a template drug

ABSTRACT

A method of forming polymers in the presence of nucleic acid using template polymerization. Also, a method of having the polymerization occur in heterophase systems. These methods can be used for the delivery of nucleic acids, for condensing the nucleic acid, for forming nucleic acid binding polymers, for forming supramolecular complexes containing nucleic acid and polymer, and for forming an interpolyelectrolyte complex.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application is a divisional of application Ser. No. 09/993,216,filed Nov. 16, 2001, Now U.S. Pat. No. 6,706,922, which is a divisionalof application Ser. No. 09/000,692, filed Dec. 30, 1997, and issued asU.S. Pat. No. 6,339,067, which is a continuation-in-part of applicationSer. No. 08/778,657, filed Jan. 3, 1997, and issued as U.S. Pat. No.6,126,964, which claims the benefit of U.S. Provisional Application No.60/009,593, filed Jan. 4, 1996.

FEDERALLY SPONSORED RESEARCH

N/A

BACKGROUND

Polymers are used for drug delivery for a variety of therapeuticpurposes. Polymers have also been used for the delivery of nucleic acids(polynucleotides and oligonucleotides) to cells for therapeutic purposesthat have been termed gene therapy or anti-sense therapy. One of theseveral methods of nucleic acid delivery to the cells is the use ofDNA-polycations complexes. It was shown that cationic proteins likehistones and protamines or synthetic polymers like polylysine,polyarginine, polyomithine, DEAE dextran, polybrene, andpolyethylenimine were effective intracellular delivery agents whilesmall polycations like spermine were ineffective. (Felgner, P. L. (1990)Advanced Drug Delivery Rev. 5, 163–187; Boussif, O., Lezoualch, F.,Zanta, M. A., Mergny, M. D., Scherman, D., Demeneix, B., & Behr, J. P.(1995) Proc. Natl. Acad. Sci. USA 92, 7297–7301) The mechanism by whichpolycations facilitate uptake of DNA is not completely understood butthe following are some important principles:

1) Polycations provide attachment of DNA to the target cell surface: Thepolymer forms a cross-bridge between the polyanionic nucleic acids andthe polyanionic surfaces of the cells. As a result the main mechanism ofDNA translocation to the intracellular space might be non-specificadsorptive endocytosis which may be more effective then liquidendocytosis or receptor-mediated endocytosis. Furthermore, polycationsare a very convenient linker for attaching specific receptors to DNA andas result, DNA-polycation complexes can be targeted to specific celltypes. (Perales, J. C., Ferkol, T. Molas, M. & Hanson, W. (1994) Eur. J.Biochem. 226, 255–266; Cotten, M., Wagner, E. & Bimstiel, M. L. (1993)Methods in Enzymology 217, 618–644; Wagner, E., Curiel, D., & Cotten, M.(1994) Advanced Drug Delivery Rev. 1 14, 113–135).

2) Polycations protect DNA in complexes against nuclease degradation(Chiou, H. C., Tangco, M. V. Levine, S. M., Robertson, D., Kormis, K.,Wu, C. H., & Wu, G. Y. (1994) Nucleic Acids Res. 22, 5439–5446). This isimportant for both extra- and intracellular preservation of DNA. Theendocytic step in the intracellular uptake of DNA-polycation complexesis suggested by results in which DNA expression is only obtained byincorporating a mild hypertonic lysis step (either glycerol or DMSO)(Lopata, M. A., D. Clevland, W., & Sollner-Webb, B. (1984) Nucleic AcidsRes. 12, 5707–5717; Golub, E. I., Kim, H. & Volsky, D. J. (1989) NucleicAcid Res. 17, 4902). Gene expression is also enabled or increased bypreventing endosome acidification with NH₄CI or chloroquine (Luthman, H.& Magnusson, G. (1983) Nucleic Acids Res. 11, 1295–1300).Polyethylenimine which facilitates gene expression without additionaltreatments probably disrupts endosomal function itself (Boussif, O.,Lezoualch, F., Zanta, M. A., Mergny, M. D., Scherman, D., Demeneix, B.,& Behr, J. P. (1995) Proc. Natl. Acad. Sci. USA 92, 7297–7301).Disruption of endosomal function has also been accomplished by linkingto the polycation endosomal-disruptive agents such as fusion peptides oradenoviruses (Zauner, W., Blaas, D., Kuechler, E., Wagner, E., (1995) J.Virology 69, 1085–1092; Wagner, E., Plank, C., Zatloukal, K., Cotten,M., & Birnstiel, M. L. (1992) Proc. Natl. Acad. Sci. 89, 7934–7938)(Fisher, K. J., & Wilson, J. M. (1994) Biochemical J. 299, 49–58).

3) Polycations generate DNA condensation: The volume which one DNAmolecule occupies in complex with polycations is drastically lower thanthe volume of a free DNA molecule. The size of DNA/polymer complex iscritical for gene delivery in vivo. In terms of intravenous injection,DNA needs to cross the endothelial barrier and reach the parenchymalcells of interest. The largest endothelia fenestrae (holes in theendothelial barrier) occurs in the liver and have an average diameter100 nm. The fenestrae size in other organs is much lower. The size ofthe DNA complexes is also important for the cellular uptake process.After binding to the target cells the DNA-polycation complex should betaken up by endocytosis. Since the endocytic vesicles have a homogenousinternal diameter of about 100 nm in hepatocytes of similar size inother cell types, the DNA complexes need to be smaller than 100 nm(Geuzze, H. J., Slot, J. W., Strous, G. J., Lodish, H. F., & Schwartz,A. L. (1982) J.Cell Biol. 92, 865–870).

Condensation of DNA

A significant number of multivalent cations with widely differentmolecular structures have been shown to induce the condensation of DNA.These include spermidine, spermine, Co(NH₃)63+, protamine, histone H1,and polylysine. (Gosule, L. C. & Schellman, J. A. (1976) Nature 259,333–335; Chattoraj, D. K., Gosule, L. C. & Schellman, J. A. (1978) J.Mol. Biol. 121, 327–337; Had, N. V., Downing, K. H. & Balhorn, R. (1993)Biochem. Biophys. Res. Commun. 193, 1347–1354; Hsiang, M. W & Cole, R.D. (1977) Proc. Natl. Acad. Sci. USA 74, 4852–4856; Haynes, M., Garret,R. A. & Gratzer, W. B. (1970) Biochemistry 9, 4410–4416; Widom, J. &Baldwin, R. L. (1980) J. Mol. Biol. 144, 431–453.). Quantitativeanalysis has shown DNA condensation to be favored when 90% or more ofthe charges along the sugar-phosphate backbone are neutralized (Wilson,R. W. & Bloomfield, V. A. (1979) Biochemistry 18, 2192–2196). Dependingupon the concentration of the DNA condensation leads to three main typesof structures:

1) In extremely dilute solution (about 1 ug/ml or below), long DNAmolecules can undergo a monomolecular collapse and form structuresdescribed as toroid.

2) In very dilute solution (about 10 ug/ml) microaggregates form withshort or long molecules and remain in suspension. Toroids, rods andsmall aggregates can be seen in such solution.

3) In dilute solution (about 1 mg/ml, large aggregates are formed thatsediment readily. (Sicorav, J.-L., Pelta, J., & Livolant, F (1994)Biophysical Journal 67, 1387–1392).

Toroids have been considered an attractive form for gene deliverybecause they have the lowest size. While the size of DNA toroidsproduced within single preparations has been shown to vary considerably,toroid size is unaffected by the length of DNA being condensed. DNAmolecules from 400 bp to genomic length produce toroids similar in size(Bloomfield, V. A. (1991) Biopolymers 31, 1471–1481). Therefore onetoroid can include from one to several DNA molecules. The kinetics ofDNA collapse by polycations which resulted in toroids is very slow. Forexample DNA condensation by Co(NH₃)6CI3 needs 2 hours at roomtemperature. (Arscott, P. G., Ma, C., & Bloomfield, V. A. (1995)Biopolymers 36, 345–364).

The mechanism of DNA condensation is not obvious. The electrostaticforces between unperturbed helices arise primarily from a counterionfluctuation mechanism requiring multivalent cations and plays the majorrole in DNA condensation. (Riemer, S. C. & Bloomfield, V. A. (1978)Biopolymers 17, 789–794; Marquet, R. & Houssier, C. (1991) J. Biomol.Struct. Dynam. 9, 159–167; Nilsson, L. G., Guldbrand, L. & NordenskjoldL. (1991) Mol. Phys. 72, 177–192). The hydration forces predominate overelectrostatic forces when the DNA helices approach closer then a fewwater diameters (Leikin, S., Parsegian, V. A., Rau, D. C. & Rand, R. P.(1993) Ann. Rev. Phys. Chem. 44, 369–395). In case of DNA-polymericpolycation interactions, DNA condensation is a more complicated processthan the case of low molecular weight polycations. Differentpolycationic proteins can generate toroid and rod formation withdifferent size DNA at a ratio of positive to negative charge of 0.4(Garciaramirez, M., & Subirana, J. A. (1994) Biopolymers 34, 285–292).It was shown by fluorescence microscopy that T4 DNA complexed withpolyarginine or histone can forms two types of structures; an elongatedstructure with a long axis length of about 350 nm (like free DNA) anddense spherical particles. (Minagawa, K., Matsuzawa, Y., Yshikawa, K.,Matsumoto, M., & Doi, M. (1991) FEBS Lett. 295, 60–67). Both forms existsimultaneously in the same solution. The reason for the co-existence ofthe two forms can be explained as an uneven distribution of thepolycation chains among the DNA molecules. The uneven distributiongenerates two thermodynamically favorable conformations. (Kabanov, A.V., & Kabanov, V. A. (1995) Bioconjugate Chem. 6, 7–20).

It was also shown that the electrophoretic mobility of DNA-polycationcomplexes can change from negative to positive in excess of polycation.It is likely that large polycations don't completely align along DNA butform polymer loops which interact with other DNA molecules. The rapidaggregation and strong intermolecular forces between different DNAmolecules may prevent the slow adjustment between helices needed to formtightly packed, orderly particles. This specification describes a newapproach, that we have termed Polynucleotide Template Polymerization,for overcoming this problem of nonspecific aggregation and largeDNA-polycation complex formation that occurs when polycation/DNAcomplexes are formed in DNA concentrations that are of practical valuefor polynucleotide transfer into cells and for gene or antisensetherapy.

SUMMARY OF INVENTION

A process for drug delivery is described in which polymerization andchemical reaction processes are induced in the presence of the drug inorder to deliver the drug or biologically active compound. Drug deliveryencompasses the delivery of a biologically active compound to a cell. By“delivering” we mean that the drug becomes associated with the cell. Thedrug can be on the membrane of the cell or inside the cytoplasm,nucleus, or other organelle of the cell. The process of delivering apolynucleotide to a cell has also been commonly termed “transfection” orthe process of “transfecting” and also it has been termed“transformation”. A biologically active compound is a compound havingthe potential to react with biological components. Pharmaceuticals,proteins, peptides and nucleic acids are examples of biologically activecompounds. The template polymer can be a polyanion such as a nucleicacid. The polynucleotide could be used to produce a change in a cellthat can be therapeutic. The delivery of polynucleotides or geneticmaterial for therapeutic purposes is commonly called “gene therapy”.

A new method is described for forming condensed nucleic acid by having achemical reaction take place in the presence of the nucleic acid. Aprocess is also described of forming in the presence of the nucleic acida polymer that has affinity to nucleic acid. Moreover, a process isdescribed of forming an interpolyelectrolyte complex containing nucleicacids by having a chemical reaction take place in the presence of thenucleic acid. In addition, the nucleic acid-binding polymer can form asa result of template polymerization. This obviously excludes theformation of polymers such as proteins or nucleic acids or otherderivatives that bind nucleic acid by Watson-Crick binding.

Previously, the occurrence of chemical reactions or the process ofpolymerization in the presence of the nucleic acid has been assiduouslyavoided when delivering nucleic acid. Perhaps, this arose out ofconcerns that the processes of chemical reactions or polymerizationwould chemically modify the nucleic acid and thereby render it notbiologically active. Surprisingly, we show that we can performpolymerizations in the presence of nucleic acids without chemicallymodifying the nucleic acid and that the nucleic acid is stillfunctional. For example, a plasmid construct containing a promoter andthe reporter gene luciferase can still express as much luciferase asnative plasmid after transfection into cells.

The process of forming a polymer in the presence of nucleic acid hasseveral advantages. As FIG. 1 illustrates, aggregation and precipitationof the nucleic acid can be avoided by having the polymerization takeplace in the presence of the nucleic acid. This newly described processenabled us to form supramolecular complexes of nucleic acid and polymerrapidly, consistently, and at very high concentrations of polynucleicacid. In fact, high concentration of the template nucleic acid favorsthis process. In contrast, the previously described process of mixing anucleic acid and an already-formed polycation (such as polylysine) hasto be done at very dilute concentrations. In addition, thepreviously-described procedure requires that the mixing, salt andionicity conditions must be carefully controlled as well. This explainswhy the use of polylysine-DNA complexes are not widely used for thetransfer of DNA into cells and is only done in a few laboratories.

The other advantage that flows from the newly described process ofhaving polymerization take place in the presence of nucleic acid is thatpolymers could form that would not be able to become associated withnucleic acids if the polymer was formed first. For example, thepolymerization process could result in a hydrophobic polymer that is notsoluble in aqueous solutions unless it is associated with nucleic acid.A hydrophobic moiety comprises a C6–C24 alkane, C6–C24 alkene, sterol,steroid, lipid, or hydrophobic hormone. Furthermore, the process ofhaving the polymerization taking place in organic solvents andheterophase systems enables more types and more defined types ofvesicles to be formed.

This process will enable supramolecular complexes to be more easilyassembled. It will also enable novel and more defined complexes to bemade. Yet another advantage that flows from this invention is thatnucleic acid/polymer complexes will be smaller. The size of DNA/polymercomplex is critical for gene delivery especially in vivo.

These processes can be used for transferring nucleic acids into cells oran organism such as for drug delivery. They may also be used foranalytical methods or the construction of new materials. They may alsobe used for preparative methods such as in the purification of nucleicacids. They are also useful for many types of recombinant DNAtechnology.

For example, they may be used to generate sequence binding molecules andprotect specific sequences from nuclease digestion. Protection ofspecific regions of DNA is useful in many applications for recombinantDNA technology.

A preferred embodiment provides a method of making a compound fordelivery to a cell, comprising: forming a polymer in the presence of abiologically active drug.

Another preferred embodiment provides a method of making a compound fordelivery to a cell, comprising: cross-linking a polymer in the presenceof a polyion, thereby forming a complex of polymer and polyion; and,delivering the complex to the cell.

Another preferred embodiment provides a method of making a compound fordelivery to a cell, comprising: modifying a molecule in the presence ofthe polyion thereby providing a deliverable polyion.

Yet, another preferred embodiment provides a method of making a compoundfor delivery to a cell, comprising: mixing a polyion with a firstpolymer and a second polymer thereby forming a deliverable complex.

Further objects, features, and advantages of the invention will beapparent from the following detailed description when taken inconjunction with the accompanying drawings.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 is a comparison of pDNA following template polymerization andcomplexation contrasted with preformed polycations binding andprecipitating.

FIG. 2 shows complexes ranging in size from 40–70 nanometers in diameterafter being dried onto carbon grids and stained with methylaminetungstate.

FIG. 3 illustrates template dependent polymerization of NLS Peptidesusing SDS-PAGE. NLS peptide multimers were observed only in thereactions when cross-linker and template (pDNA) were present together(panel A, lanes 6 and 7), NLS peptides migrated at positionscorrecponding to monomers when treated with (dithiothreitol) DTT priorto SDS-PAGE (panel B, lanes 6 and 7).

FIG. 4 illustrates the relationship between turbidity (an indication ofaggregation) and the molar charge ratio of polylysine (PLL):plasmid DNAwithout and with 100 mM NaCl and without and without templatepolymerization/caging (indicated by +DTBP(dimethyl-3,3′-dithiobispropionimidate)).

FIG. 5 illustrates the ability of dextran sulfate (DS) to enableethidium bromide to interact with plasmid DNA that has been complexedwith varying ratios of PLL/DNA (molar ratio of lysine residue to DNAbase) (numbers in the legends) with or without the addition ofdimethyl-3,3′-dithiobispropionimidate (DTBP).

FIG. 6 illustrates the effect of varying the histone/DNA ratio on thesizes of histone/DNA complexes with the addition ofdimethyl-3,3′-dithiobispropionimidate (DTBP).

DETAILED DESCRIPTION

1. Drug Delivery

A process for drug delivery is described in which polymerization andchemical reaction processes take place in the presence of the drug inorder to deliver the drug. The polymer is formed from a variety ofmonomers in the presence of the drug and then the mixture is delivered.The mixture could undergo further purification or preparative methods.Drug delivery encompasses the delivery of a biologically active compoundto a cell. This can be accomplished with prokaryotic or eukaryotic cell.It includes mammalian cells that are either outside or within anorganism. It also includes the administration of the drug to the wholeorganism by standard routes such as intravenous, intra-arterial,intra-bile duct, intramuscular, subcutaneous, intraperitoneal, or directinjections into tissues such as the liver, brain, kidneys, heart, eyes,lymph nodes, bone, gastrointestinal tract. It also includes deliveryinto vessels such as blood, lymphatic, biliary, renal, or brainventricles.

In one preferred embodiment, this process is used to deliver nucleicacids. The process of delivering nucleic acids means exposing the cellto the polynucleic in the presence of the delivery system. Cellsindicate both prokaryotes and eukaryotes. The cell is located in aliving organism and exposing is accomplished by administering thenucleic acid and the delivery system to the organism. It also meansmixing the nucleic acids with cells in culture or administering thenucleic acids to a whole organism. Delivering nucleic acids encompassestransfecting a cell with a nucleic acid. These delivery processesinclude standard injection methods such as intramuscular, subcutaneous,intraperitoneal, intravenous, and intra-arterial. It also includesinjections into any vessel such as the bile duct and injections into anytissue such as liver, kidney, brain, thymus, heart, eye, or skin.

Drugs, pharmaceuticals, proteins, peptides and nucleic acids arebiologically active compounds. The drug can be either the templatepolymer or the daughter polymer. In the preferred embodiment, thetemplate polymer is a polyion, a macromolecule carrying a string ofcharges, such as a nucleic acid which would be termed a polyanionbecause of its average negative charge. The term “nucleic acid” is aterm of art that refers to a string of at least two base-sugar-phosphatecombinations. Nucleotides are the monomeric units of nucleic acidpolymers. The term includes deoxyribonucleic acid (DNA) and ribonucleicacid (RNA) in the form of an oligonucleotide, messenger RNA, anti-sense,plasmid DNA, parts of a plasmid DNA or genetic material derived from avirus. The term “nucleic acid” includes both oligonucleic acids andpolynucleic acids. Polynucleic acids are distinguished from oligonucleicacid by containing more than 120 monomeric units. In the case of thetransfer of nucleic acids into cells, the nucleic acid is the template.

The nucleic acid (polynucleotide) could also be used to produce a changein a cell that can be therapeutic. The delivery of polynucleotides orgenetic material for therapeutic purposes is commonly called “genetherapy”. The delivered polynucleotide could produce a therapeuticprotein such as a hormone, cytokine, or growth factor. For example, thepolynucleotide in the form of a plasmid DNA could produce the humangrowth hormone. The polynucleotide could produce an enzyme that isdeficient or defective in patients with an inborn error of metabolism.For example, a plasmid DNA could produce phenylalanine hydroxylase whichwould be therapeutic in patients with phenylketonuria. Furthermore, thepolynucleotide could supply an anti-sense that would be therapeutic inpatients with a tumor, cancer, or infection. For example, thepolynucleotide could be a DNA that is transcribed into an anti-sensemolecule.

2. Formation of Polymers

A polymer is a molecule built up by repetitive bonding together ofsmaller units called monomers. In this application the term polymerincludes both oligomers which have two to ˜80 monomers and polymershaving more than 80 monomers. The polymer can be linear, branchednetwork, star, comb, or ladder types of polymer. The polymer can be ahomopolymer in which a single monomer is used or can be copolymer inwhich two or more monomers are used. Types of copolymers includealternating, random, block and graft.

Associated with the polymer in a preferred embodiment is a stericstabilizer which is a long chain hydrophilic group that preventsaggregation of final polymer by sterically hindering particle toparticle electrostatic interactions. Examples include: alkyl groups, PEGchains, polysaccharides, hydrogen molecules, alkyl amines.

To those skilled in the art of polymerization, there are severalcategories of polymerization processes that can be utilized in thedescribed process. The polymerization can be chain or step. Thisclassification description is more often used that the previousterminology of addition and condensation polymer. “Most step-reactionpolymerizations are condensation processes and most chain-reactionpolymerizations are addition processes” (M. P. Stevens PolymerChemistry: An Introduction New York Oxford University Press 1990).

2A. Step Polymerization

In step polymerization, the polymerization occurs in a stepwise fashion.Polymer growth occurs by reaction between monomers, oligomers andpolymers. No initiator is needed since there is the same reactionthroughout and there is no termination step so that the end groups arestill reactive. The polymerization rate decreases as the functionalgroups are consumed.

Typically, step polymerization is done either of two different ways. Oneway, the monomer has both reactive functional groups (A and B) in thesame molecule so that

-   -   A-B yields -[A-B]

Or the other approach is to have two difunctional monomers.

-   -   A-A+B-B yields -[A-A-B-B]

Generally, these reactions can involve acylation or alkylation.Acylation is defined as the introduction of an acyl group (—COR) onto amolecule. Alkylation is defined as the introduction of an alkyl grouponto a molecule.

If functional group A is an amine then B can be (but not restricted to)an isothiocyanate, isocyanate, acyl azide, N-hydroxysuccinimide,sulfonyl chloride, aldehyde (including formaldehyde and glutaraldehyde),epoxide, carbonate, imidoester, carboxylate, or alkylphosphate,arylhalides (difluoro-dinitrobenzene) or succinic anhyride. In otherterms when function A is an amine then function B can be acylating oralkylating agent.

If functional group A is a sulfhydryl then function B can be (but notrestricted to) an iodoacetyl derivative, maleimide, aziridinederivative, acryloyl derivative, fluorobenzene derivatives, or disulfidederivative (such as a pyridyl disulfide or 5-thio-2-nitrobenzoicacid{TNB} derivatives).

If functional group A is carboxylate then function B can be (but notrestricted to) a diazoacetate or an amine in which carbonyldiimidazoleor carbodiimide is used.

If functional group A is an hydroxyl then function B can be (but notrestricted to) an epoxide, oxirane, or an amine in whichcarbonyldiimidazole or carbodiimide or N,N′-disuccinimidyl carbonate, orN-hydroxysuccinimidyl chloroformate is used.

If functional group A is an aldehyde or ketone then function B can be(but not restricted to) an hydrazine, hydrazide derivative, aldehyde (toform a Schiff Base that may or may not be reduced by reducing agentssuch as NaCNBH₃).

Yet another approach is to have one difunctional monomer so that

-   -   A-A plus another agent yields -[A-A]-.

If function A is a sulfhydryl group then it can be converted todisulfide bonds by oxidizing agents such as iodine (I₂) or NaIO₄ (sodiumperiodate), or oxygen (O₂). Function A can also be an amine that isconverted to a sulfhydryl group by reaction with 2-Iminothiolate(Traut's reagent) which then undergoes oxidation and disulfideformation. Disulfide derivatives (such as a pyridyl disulfide or5-thio-2-nitrobenzoic acid{TNB} derivatives) can also be used tocatalyze disulfide bond formation.

Functional group A or B in any of the above examples could also be aphotoreactive group such as aryl azides, halogenated aryl azides, diazo,benzophenones, alkynes or diazirine derivatives.

Reactions of the amine, sulfhydryl, carboxylate groups yield chemicalbonds that are described as amide, amidine, disulfide, ethers, esters,isothiourea, isourea, sulfonamide, carbamate, carbon-nitrogen doublebond (enamine or imine) alkylamine bond (secondary amine),carbon-nitrogen single bonds in which the carbon contains a hydroxylgroup, thio-ether, diol, hydrazone, diazo, or sulfone.

2B. Chain Polymerization

In chain-reaction polymerization growth of the polymer occurs bysuccessive addition of monomer units to limited number of growingchains. The initiation and propagation mechanisms are different andthere is usually a chain-terminating step. The polymerization rateremains constant until the monomer is depleted.

Monomers containing vinyl, acrylate, methacrylate, acrylamide,methaacrylamide groups can undergo chain reaction which can be radical,anionic , or cationic. Chain polymerization can also be accomplished bycycle or ring opening polymerization. Several different types of freeradical initiatiors could be used that include peroxides, hydroxyperoxides, and azo compounds such as 2,2′-Azobis(-amidinopropane)dihydrochloride (AAP).

3. Types of Monomers

A wide variety of monomers can be used in the polymerization processes.These include positive charged organic monomers such as amines, imidine,guanidine, imine, hydroxylamine, hydrozyine, heterocycles (likeimidazole, pyridine, morpholine, pyrimidine, or pyrene. The amines couldbe pH-sensitive in that the pKa of the amine is within the physiologicrange of 4 to 8. Specific amines include spermine, spermidine,N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD), and3,3′-Diamino-N,N-dimethyldipropylammonium bromide (Compound 9).

-   1. Monomers can also be oligopeptides, polypeptides or proteins    (produced synthetically or in an organism). These oligopeptides can    be a NLS peptide which corresponds to the 12 amino acid nuclear    localizing sequence of SV40 T antigen, fusion peptides (derived from    viruses), endosomolytic peptides and amphipathic peptides.    Amphipathic compounds have both hydrophilic (water-soluble) and    hydrophobic (water-insoluble) parts. The amphipathic compound can be    cationic or incorporated into a liposome that is positively-charged    (cationic) or non-cationic which means neutral, or    negatively-charged (anionic). Proteins such as histone H1 can be    used. Proteins that bind DNA at sequence-specific sequences such as    Gal4 protein could also be used.

Monomers can also be hydrophobic, hydrophilic or amphipathic. Examplesof amphipathic compounds include but are not restricted to3,3′-diamine-N-(7-octene)-N-methyldipropylammonium bromide (Compound 7),N,N′-Dinonacrylate-N,N,N′,N′-tetramethylpropanediammonium bromide(Compound 10),N,N′,N″-Trinonacrylate-N,N,N′,N′,N″-pentamethyldiethylentriammoniumbromide (Compound 11) and amphipathic peptides such asCKLLKKLLKLWKKLLKKLKC.

Monomers can also be intercalating agents such as acridine, thiazoleorgange, or ethidium bromide.

4. Other Components of the Monomers and Polymers

The polymers have other groups that increase their utility. These groupscan be incorporated into monomers prior to polymer formation of attachedto the polymer after its formation. These groups include:

a. Targeting Groups—These groups are used for targeting the polymer-drugor polymer-nucleic acid complexes to specific cells or tissues. Examplesof such targeting agents include agents that target to theasialoglycoprotein receptor by using asiologlycoproteins or galactoseresidues. Other proteins such as insulin, EGF, or transferrin can beused for targeting. Peptides that include the RGD sequence can be usedto target many cells. Chemical groups that react with sulfhydryl ordisulfide groups on cells can also be used to target many types ofcells. Folate and other vitamins can also be used for targeting. Othertargeting groups include molecules that interact with membranes such asfatty acids, cholesterol, dansyl compounds, and amphotericinderivatives.

After interaction of the supramolecular complexes with the cell, othertargeting groups can be used to increase the delivery of the drug ornucleic acid to certain parts of the cell. For example, agents can beused to disrupt endosomes and a nuclear localizing signal (NLS) can beused to target the nucleus.

A variety of ligands have been used to target drugs and genes to cellsand to specific cellular receptors. The ligand may seek a target withinthe cell membrane, on the cell membrane or near a cell. Binding ofligands to receptors typically initiates endocytosis. Ligands could alsobe used for DNA delivery that bind to receptors that are notendocytosed. For example peptides containing RGD peptide sequence thatbind integrin receptor could be used. In addition viral proteins couldbe used to bind cells. Lipids and steroids could be used to directlyinsert into cellular membranes.

b. Reporter Groups—Reporter or marker groups are molecules that can beeasily detected. Typically they are fluorescent compounds such asfluorescein, rhodamine, texas red, cy 5, or dansyl compounds. They canbe molecules that can be detected by UV or visible spectroscopy or byantibody interactions or by electron spin resonance. Biotin is anotherreporter molecule that can be detected by labeled avidin. Biotin couldalso be used to attach targeting groups.

c. Cleavable Groups—The polymers can contain cleavable groups within thetemplate binding part or between the template binding part and thetargeting or reporter molecules. When within the template binding part,breakage of the cleavable groups leads to reduced interaction of thetemplate and daughter polymers. When attached to the targeting group,cleavage leads to reduce interaction between the template and thereceptor for the targeting group. Cleavable groups include but are notrestricted to disulfide bonds, diols, diazo bonds, ester bonds andsulfone bonds.

5. Template Polymerization

Template polymerization has been defined as the following (van deGrampel, H. T., Tan, Y. Y. and Challa, G. Macromolecules 23, 5209–5216,1990):

“Template polymerizations can be defined as polymerizations in whichpolymer chains are able to grow along template macromolecules for thegreater part of their lifetime. Such a mode of propagation can beachieved through the existence of cooperative interactions between thegrowing chain and the template chain and usually leads to the formationof an interpolymer complex. In general, a well-chosen template is ableto affect the rate of polymerization as well as the molecular weight andmicrostructure of the formed polymer (daughter polymer). The concepts oftemplate polymerization were described by Ballard and Bamford with thering opening polymerization of the N-carboxyanhydride ofDL-phenylalanine on a polysarcosine template. Since then, many othersystems involving radical and nonradical initiation of vinyl monomershave been studied in which one or more template effects, arising fromthis peculiar propagation mode, were identified. A number ofradical-initiated template polymerizations have been studied, employingwater as solvent”.

The main features of template polymerization are:

1. Complex formation takes place between polymers

2. The rate of polymerization increases as the concentration of templateincreases. (Fujimori, K., (1979) Makromol. Chem. 180, 1743)

3. The structure and conformational features of the template arereflected in the corresponding daughter polymer.

In template polymerization, propagation of new polymer chain occurspredominantly along the template, a macromolecular chain, throughspecific cooperative interaction. The nature of interaction can beelectrostatic, H—bonding, charge-transfer, and Van der Waals forces incombination with steriochemical matching. The presence of templateusually affects various polymerization characteristics as well as themicrostructure of the polymer formed. The mechanism of templatepolymerization depends on the degree of monomer adsorption. Two extremecases can be discerned: the adsorption equilibrium coefficient formonomer, K_(M)=·(type 1) and K_(M)=0 (type 2). In type 1 (“zip”reaction) monomer is fully adsorbed onto all template sites and thepolymerization occurs only on template. As the K_(M) constant becomessmaller, template propagation increasingly proceeds via reactionmonomers from the surrounding solution at the expense of reaction withadjacently adsorbed monomer. When K_(M)=0 (type 2) only non-adsorbedmonomer is present and the template macromolecules are completelysolvated by solvents instead of the monomers. A prerequisite fortemplate propagation under this condition is the growing daughteroligomer, created in bulk solution, that then complexes with template.(“pick-up” reaction). The chain length below which no complexation takesplace (critical chain length) is important for magnitude of the templateeffect. In fact, there is no sharp border between type 1 and type 2polymerization's.

Several processes for using template polymerization for drug deliveryare described. The daughter polymer could be the drug. In a preferredembodiment, the template is the drug (defined to includepharmaceuticals, therapeutic agents or biologically active substances).The process of using template polymerization for drug delivery comprisesmixing the template with monomers and having a daughter polymer formingfrom the monomers. The mixture of template polymer and daughter polymeris then administered to a cell by putting the mixture in contact with acell or near a cell. The mixture of template and daughter polymer couldalso be placed in a pharmaceutical formulation and vial for delivery toan animal. The template polymer could be a polyanion such as nucleicacid including DNA, RNA or an antisense sequence. The DNA can produce atherapeutic agent such as a therapeutic protein or anti-sense RNA.

In a preferred embodiment, targeting groups could be added during theinitial template polymerization stage or during subsequentpolymerization steps. In addition, after template polymerization,networks or additional networks can be added to the polymer. These couldbe used to cross-link the polymers. For example, the polymer could becross-linked to put the template into a “cage”. Cross-linking is thelinking of two moieties of a polymer to one another using bifunctionalchemical linker. One result is that the polymer, as a network, becomesstronger and more resistant to being dissolved. Covalent linkingbifunctional linkers may be homobifunctional (which involves the samechemical reaction for linking both moieties) or heterobifunctional(involves two different reactions allowing linkage of differentfunctional groups). By cross-linking, a cage may be formed around ornear the polyion creating a complex of polyion and polymer.Cross-linking the polymer protects the polyion from being destroyed byenzymes and other degrading substrates. For example: If the polyion isDNA, the cross-linked or caging polymer protects DNA from DNases.

In a preferred embodiment, stable caged polyion particles still bear anet positive charge. However, it is desirable to recharge it so it wouldinteract less with negatively-charged polymers and particles in vivo.Recharging is switching the net polyion particle charge to an oppositecharge.

Complexes may be formed and continue to function in a solution ofchangeable tonicity, which means that the solution can be hypotonic,hypertonic or normal tonicity. Hypotonic means any solution which has alower osmotic pressure than another solution (that is, has a lowerconcentration of solutes than another solution). A hypotonic solution isthe opposite of a hypertonic solution. Normal tonicity in the preferredembodiments is the tonicity of human body fluids, specifically blood.

6. Homophase and Heterophase Polymerization

The chemical reaction and polymerization processes can take place inhomophase systems in which the monomer and nucleic acid are in the samesolution. This solution can be water, alcohol, chloroform, esters,organic solvents, or polar aprotic solvents such as DMF or DMSO ordioxane. They can be mixtures of aqueous and organic solvents.

The chemical reaction and polymerization processes can take place inheterophase systems in which the nucleic acid is in one phase and themonomer is in another phase. Such heterophase systems can be “oil inwater” and also “water in oil” where oil is defined as a solvent thathas low solubility in water. This approach could enable the formation ofmicellar-like structures that have the hydrophobic parts of thepolynucleotide in the inside of a vesicle and the hydrophilic parts onthe outside, or vice-versa. The polymerization reaction can be performedin both direct (oil-in-water) and inverse (water-in-oil) emulsions. Thisapproach allows the use of hydrophobic or amphipathic monomers(Blackley, D. C. Emulsion Polymerization, London: Appl. Sci., 1975).Heterophase polymerization enables vesicles, particles, orsupramolecular complexes to be produced in which the nucleic acid is onthe surface of polymer micelles or the nucleic acid is inside ofmonolayer inverse polymer micelles. In the last case different lipidscan be used for external layer formation. Inverse phase emulsion can beprepared so that in average only one molecules of biopolymer will bepresent in every water drop. (Martinek, K., Levashov, A. V., Klyachko,N., Khmelnitski, Y. L., & Berezin, I. V. (1986) Eur. J. Biochem. 155,453–468).

7. Supramolecular Complexes

A supramolecular complex is a structure that contains two or moredifferent molecules that are not covalently bound. Supramolecularcomplexes can be used for drug delivery and for other purposes such asfor preparative or analytical methods or the construction of newmaterials. We describe a new method for forming a supramolecular complexcontaining nucleic acid and a polymer in which the polymer is formed inthe presence of the nucleic acid.

The supramolecular complex can contain other components in addition tothe nucleic acid and polymer. It can contain another polymer that isalready formed. This already formed polymer can bind the nucleic acid orthe daughter polymer. The additional component can be a protein. Thisprotein can be cationic and contain positive charges that enables it tobind nucleic acid. Such cationic proteins could be histone, polylysine,or protamine. The supramolecular complex could also contain targetinggroups.

A supramolecular complex formed in this fashion could containamphipathic compounds that could be part of liposomes, micelles, orinverse micelles. Liposomes are microscopic vesicles that containamphipathic molecules that contain both hydrophobic and hydrophilicdomains. Liposomes can contain an aqueous volume that is entirelyenclosed by a membrane composed of lipid molecules (usuallyphospholipids) (R. C. New, p. 1, chapter 1, “Introduction” inLiposomes:A Practical Approach, ed. R.C. New IRL Press at OxfordUniversity Press, Oxford 1990). Micelles and inverse micelles aremicroscopic vesicles that contain amphipathic molecules but do notcontain an aqueous volume that is entirely enclosed by a membrane. Inmicelles the hydrophilic part of the amphipathic compound is on theoutside (on the surface of the vesicle) whereas in inverse micelles thehydrophobic part of the amphipathic compound is on the outside.

8. Condensed Nucleic Acids

A method of condensing nucleic acid is defined as decreasing the linearlength of the nucleic acid. Condensing nucleic acid also meanscompacting nucleic, acid. Condensing nucleic acid also means decreasingthe volume which the nucleic acid molecule occupies. A example ofcondensing nucleic acid is the condensation of DNA that occurs in cells.The DNA from a human cell is approximately one meter in length but iscondensed to fit in a cell nucleus that has a diameter of approximately10 microns. The cells condense (or compacts) DNA by a series ofpackaging mechanisms involving the histones and other chromosomalproteins to form nucleosomes and chromatin. The DNA within thesestructures are rendered partially resistant to nuclease (DNase) action.The condensed structures can also be seen on electron microscopy.

The process of condensing nucleic acid can be used for transferringnucleic acids into cells or an organism such as for drug delivery. Itcould also be used for prepartive or analytical methods or theconstruction of new materials.

We describe a new method for forming condensed nucleic acid by having achemical reaction take place in the presence of the nucleic acid. Achemical reaction is defined as a molecular change in the participantatoms or molecules involved in the reaction. An example of a molecularchange would be the breaking and forming of covalent bonds ofparticipant compounds. Covalent bonds are defined as havingshared-electron bonds such as those found, in carbon-carbon,carbon-nitrogen, carbon hydrogen, carbon-oxygen, carbon-sulfur,carbon-halogen, nitrogen-hydrogen, oxygen-hydrogen, oxygen-oxygen andsulfur-oxygen bonds. The chemical reaction(s) could result in a polymerbeing formed. The polymerization process could take place by the processof template polymerization. A supramolecular complex could form as aresult of this process.

In a preferred embodiment, one method utilizes covalently attachingcompounds to polyions such as genes for enhancing the cellular transportof the polyion while maintaining its functionality. A patent applicationentitled: A Method For Covalent Attachment Of Compounds To Genes, Ser.No. 08/990,015, filed Dec. 12, 1997 describing methods of covalentlyattaching compounds as well as structures used therein is incorporateherein by reference. Although the cited application teaches theattachment of compounds to genes, the methods and structures may beapplied to attaching molecules to a polymer as discussed in the presentspecification and the term polymer is not limited to nucleic acids.

Signals that enhance release from intracellular compartments (releasingsignals) can cause polyion release from intracellular compartments suchas endosomes (early and late), lysosomes, phagosomes, vesicle,endoplasmic reticulum, golgi apparatus, trans golgi network TGN), andsarcoplasmic reticulum. Release includes movement out of anintracellular compartment into cytoplasm or into an organelle such asthe nucleus. Releasing signals include chemicals such as chloroquine,bafilomycin or Brefeldin A1 and the ER-retaining signal (KDEL sequence),viral components such as influenza virus hemagglutinin subunit HA-2peptides and other types of amphipathic peptides.

Nuclear localizing signals enhance the entry of a polyion into thenucleus or directs the gene into the proximity of the nucleus. Suchnuclear transport signals can be a protein or a peptide such as the SV40large T ag NLS or the nucleoplasmin NLS.

9. A Method for Forming A Polymer That Binds Nucleic Acids

We describe a process of forming in the presence of the nucleic acid apolymer that has affinity to nucleic acid. This excludes the process offorming polymers that are proteins or nucleic acids. It also excludespolymers that bind the nucleic acid by Watson-Crick binding.Watson-Crick binding is defined as the normal base-pairing arrangementin which guanine base pairs with cytosine base and in which theadenosine base pairs with thymine bases. Affinity indicates that thepolymer is attracted to nucleic acid and remains bound to it bynon-covalent forces (such as van der waal, hydrogen bonds, and ionicbonds) under either physiologic or non-physiologic conditions.

The process of forming a polymer in the presence of the nucleic acid canbe used for transferring nucleic acids into cells or an organism such asfor drug delivery. It could also be used for preparative or analyticalmethods or the construction of new materials.

The nucleic acid-binding polymer can form as a result of templatepolymerization.

We also describe a process of forming an interpolyelectrolyte complexcontaining nucleic acids by having a chemical reaction take place in thepresence of the nucleic acid. An interpolyelectrolyte complex is definedas a mixture of two polymers with opposite charges. In this situationthe nucleic acid is a polyanion and the formed polymer is a polycation.

Definitions of Compounds used in Preferred Embodiments

Orthogonal—Refers to a protective (protecting) group that can beselectively removed in the presence of other protective groups containedon the molecule of interest.

Monovalent—refers to an ionic species possessing 1 charge.

Protective group—A chemical group that is temporarily bound tofunctionalities within a multifunctional compound that allows selectivereactions to take place at other sites within the compound. Commonprotective groups include, but are not limited to carbamates, amides,and N-alkyl groups.

Functionality—Refers to general classes of organic compounds such as:alcohols, amines, carbonyls, carboxyls, and thiols.

EXAMPLES

Overview of Experimental Design

The following examples show that polymerization can take place in thepresence of DNA. Since the central feature of these polymers is theirability to bind DNA, we selected a relatively simple assay to detect theformation of such polymers and that is agarose gel electrophoresis withethidium bromide staining of DNA. A strong DNA-binding polymer retards(or slows) the migration of the DNA in the gel. In the experimentalsamples where the DNA is already present during the polymerization(reaction) process, the sample is simply loaded onto the agarose gel. Inthe control samples where DNA is not present during the reactionprocess, the DNA is added after the reaction. This approach is also apowerful method to determine whether any polymer is formed by a templatepolymerization process. That is, if the polymer only forms when thetemplate DNA is present and not when the template DNA is absent thenthis is definitive proof of template polymerization. The initial resultswith agarose gel electrophoresis are followed up with more sophisticatedassay for polymers and particles that include gel filtration (sizeexclusion) chromatography, transmission electron microscopy, andparticle sizing by dynamic light scattering.

The process of polymerization in the presence of nucleic acids can beused to transfer and express genes in cells. Besides showing the utilityof this process, it also indicates that the chemical reactions were notchemically modifying or destroying the nucleic acid. A another approachwas also used to detect nucleic acid damage. We incorporated disulfidebonds into the polymers and then broke the polymers down by addingdithiothreitol (DTT also known as Cleland's reagent) which reduces thedisulfide bonds. After the breakdown of the polymers the nucleic acid,DNA (that was within the polymer particles) was transfected into cellsusing another transfection method (with a cationic lipid). Expressionwas the same as the native DNA. Expression is a very sensitive indicatorof any destruction or modification along the entire length of thereporter (luciferase) gene and promoter. These polymers were designedwith disulfide bonds so that they could more easily be broken downinside cells.

Example 1

Step polymerization with DNA as a template was performed using thepolyamine N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD) anddithiobis(succinimidylpropionate) (DSP). This template polymerizationwas done using two different monomer species together in which each ofthe species possessed at least two reactive ends to propagate a growingchain. Using a bifunctional amine with affinity to plasmid DNA as amonomer and bifunctional aminoreactive cross-linker as a co-monomer, wedemonstrated that 1) it is possible to obtain DNA-bound polyamide as aresult of such polymerization, and 2) the resulting polymer can condensetemplate DNA into compact structures.

Methods:

The following amine was used as monomers:

-   1. N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD, Aldrich,    Milwaukee, Wis.)

The following cross-linker was used as a co-monomer:

-   1. Dithiobis(succinimidylpropionate), (DSP, S-S cleavable bis    succinimide ester, Pierce, Rockford, Ill.)

Optimized reaction conditions with AEPD/DSP were as follows. Plasmid DNA(pCIluc, 50 mg) and AEPD (10 mg) were mixed in 50 ml of buffer solution(0.1 M HEPES, 1 mM EDTA, pH 7.4). After 5 min DSP (60 mg in 1.5 ml ofdimethylformamide) was added. After mixing, the reaction was left for 1hour in the dark at room temperature. Finally, reaction mixture wasdialysed against water or desired buffer solution in microdialysis cellwith a molecular weight cut-off of 1,000 (Rainin, Ridgefield, N.J.).

The pCILuc plasmid expresses a cytoplasmic luciferase from the humanimmediately early cytomegaloviral (CMV) promoter. It was constructed byinserting the cytoplasmic luciferase cDNA into the pCI (Promega Corp.,Madison, Wis.) CMV expression vector. Specifically, a Nihau/EcoRIrestriction digestion fragment containing the cytoplasmic luciferasecDNA was obtained from pSPLuc (Promega Corp.) and inserted into pCI pDNAthat was digested with NheI and EcoRI. Plasmid DNA was purified usingthe Qiagen (Chatsworth, Calif.) plasmid purification system (alkalinelysis followed by anion exchange chromatography).

Agarose gel electrophoresis and ethidium bromide staining of the DNA wasdone using standard techniques (Sambrook, J., Fritsch, E. F., andManiatis, T. (1989) in Molecular Cloning Cold Spring Harbor LaboratoryPress, Cold Spring Harbor, N.Y.).

Standard gel filtration (size-exclusion) chromatography was performed todetermine the size of the polymers that formed in the presence andabsence of DNA. Since the DNA strongly bound the polymer, it wasnecessary to first remove the DNA. This was accomplished by vigorousDNase digestion. Samples of DNA/AEPD/DSP reaction mixture (50 ug totalDNA, pCIluc) were supplemented with 5 M NaCl solution up to 0.5 M NaCl.DNase I (Sigma) was added to the mixture (0.06 U/ug DNA). DNasedigestion was carried out in the buffer containing 10 mM Tris, 10 mMMgCl2, 1 mM CaCl₂, pH 7.0, for 4 hrs at 37° C. After this reaction, themixture was centrifuged at 12,000 rpm for 5 min and applied on theSephadex G-75 (Sigma) column (0.8×20 cm) equilibrated with 20 mM HEPES,0.5 NaCl, pH 7.4. Fractions (0.5 ml) were analyzed for OD 260.

Transmission electron microscopy of the formed complexes using standardnegative staining procedures on coated grids. After the samples werestained with methylamine tungstate (BioRAD), the grids were examinedusing a Jeol 100CX transmission electron microscope.

The preparation for light scattering was prepared essentially with thesame DNA/AEPD/DSP ratios as for EM (see optimized AEPD/DSP) but with 3mg of DNA (pCIluc). DNA/AEPD mixture was incubated with occasionalvortexing for 10 min at room temperature before addition of DSP. Thesample was centrifuged at 12,000 rpm for 5 min and passed through 0.2 umpolycarbonate filter (Poretics Corp., Livermore, Calif.) and analyzedusing Particle Size Analyser equipped with 15 M argon laser (BrookhavenInstruments, Inc.).

Results:

Agarose gel electrophoresis of the final experimental complexes (formedby reacting AEPD and DSP in the presence of DNA) demonstratedcharacteristic gel retardation of plasmid DNA in the gel in which thecomplexed plasmid DNA migrated more slowly than the original plasmidDNA. In addition there was some DNA material in the starting well. Thefinal complexes were also treated with 25 mM dithiothreithol (DTT)(Fisher, Itasca Ill.) for 30 minutes at 37° C. to cleave the disulfidebonds within the polymer (part of the DSP co-monomer). The DTT treatmentreversed the electrophoretic pattern back to that of the native plasmidDNA and no retarded DNA material was present. This indicates that theretarded pattern was due to the polymer complexing with the DNA. It alsoindicates that the monomers or polymer did not react with the DNA.Transfection of the DNA (after DTT treatment) into cells in cultureusing a commercial transfection reagent (LT-1, Mirus, Madison, Wis.)resulted in as much luciferase expression as native DNA. This alsoindicates that the DNA was not chemically modified.

A control sample contained AEPD and DSP at the same concentrations butplasmid DNA was omitted during the reaction. The DNA was added after thereaction was completed. Agarose gel electrophoresis showed much lessretardation of the DNA than the above experimental sample. Thisindicates that polymerization did not occur in the control sample andthat the polymerization observed in the experimental sample occurred bytemplate polymerization.

Further studies were performed to determine the size of the polymer thatformed in the presence of the DNA. This was accomplished by firstdigesting the DNA exhaustively with DNase and then running the remainingpolymer through a size-exclusion column. Gel filtration of the complex'sexhaustive DNase lysate in 0.5 M NaCl demonstrated formation of theproduct with apparent molecular weight of >3,000 Da. The control sample(DNA added after reaction of AEPD and DSP) did not contain any largepolymer of this molecular weight. This indicates that the polymer thatformed in the presence of DNA occurred by template polymerization.

Physical methods were employed to determine directly the size and shapeof the polymer/DNA complexes. Transmission electron microscopy of theexperimental complexes (formed by reacting AEPD and DSP in the presenceof DNA) revealed formation of spherical structures with 40–50 nm indiameter (individual and aggregated) (FIG. 2). Dynamic light scatteringof the same preparation yielded average particle size of 80 nm. Theseresults are consistent with the ability for the particles to passthrough 0.2 micron filters. The control samples (DNA added afterreaction of AEPD and DSP) did not contain any particles on electronmicroscopy or particle sizing.

Findings:

1. The polyamine was co-polymerized with DSP to form a polymer in thepresence of DNA and this polymer was bound to the DNA.

2. The polymer formed by a process of template polymerization.

3. The polymer condensed the DNA to form particles less than 80 nm indiameter.

4. The DNA was not chemically modified by the polymerization process andwas still able to express luciferase after transfection into cells inculture.

Example 2

Step polymerization with DNA as a template was performed using thepolyamine N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD) as in Example1 above except DPBP was used as the co-monomer.

Methods:

The following amine was used as monomers:

1. N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD, Aldrich, Milwaukee,Wis.)

The following cross-linker was used as a co-monomer:

-   1. Dimethyl-3,3′-dithiopbispropionimidate (DPBP, S-S cleavable    bisimido ester, Pierce)

Optimized reaction conditions with AEPD/DPBP were as follows. PlasmidDNA (pCIluc, 50 mg) and AEPD (24 mg) were mixed in 150 ml of buffersolution (20 mM HEPES, 1 mM EDTA, pH 7.4). After 5 min DPBP (155 mg in 5ml of methanol) was added. After mixing, the reaction was left for 1hour in the dark at room temperature.

Results:

Unlike the bissuccinimidate reaction (example 1), diimidoestercross-linking (used in this example) preserves positive charges ofaminogroups by converting them into amidines. Therefore, extremelypositively charged polymer was formed as a result of this reaction whichcaused complete DNA retardation on agarose gels. DNA addition to thereaction mixture after the reaction between amine and cross-linker didnot induce DNA retardation on the gel. Treatment of retarded complexeswith DTT resulted in restoration of the native plasmid electrophoreticpattern.

Findings:

1. Step polymerization of AEPD and DPBP occurred in the presence of DNAand resulted in a polymer that bound DNA very strongly.

2. The polymer formed by template polymerization.

3. The DNA was not chemically modified and could be recovered intactafter DTT treatment.

Example 3

Step polymerization with DNA as a template was performed using thepolyamine N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD) as in Example1 above except 2-iminothiolane (Traut's reagent) was used as theco-monomer. This is an example of ring opening of the 2-iminothilane andthen oxidation of SH groups that form as a result of the ring opening.

Methods:

The following amine was used as monomers:

-   1. N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD, Aldrich,    Milwaukee, Wis.)

The following cross-linker was used as a co-monomer:

-   1. 2-iminothiolane (thiol/amino-linking heterobifunctional agent,    Pierce)

Optimized reaction conditions with AEPD/2-iminothiolane were as follows.Plasmid DNA (pCILuc, 50 mg), AEPD (1 mM) and iminothiolane (4 mM) weremixed in 450 ml of buffer solution (20 mM HEPES, 1 mM EDTA, pH 8.0).After 30 min 5 ml of iodine solution (40 mM in ethanol) were added.Reaction was allowed to stand for 1.5 h in the dark at room temperature.

Results:

Generally, the above procedure is two-step polymerization with reactivemonomer formation. 2-iminothiolane forms bisthiol AEPD derivative on theDNA which can be further polymerized by oxidizing SH groups withmolecular iodine. Results for DNA gel retardation and DTT treatment arebasically the same as for AEPD/DPBP pair. Under conditions indicatedabove DNA and AEPD/2-iminothiolane polymer form truly soluble complexcompletely retarded in agarose gel.

The DNA in the control sample (DNA added after the polymer reaction) wasnot retarded on gel electrophoresis.

Findings:

1. Ring opening and two-step polymerization processes can be used forforming template polymers that bind to DNA.

2. Polyamines can be polymerized in the presence of DNA using theconversion amines to SH groups with subsequent oxidation reactions.

Example 4

Similar results were obtained when spermine was used instead of AEPD asin Example 1. Plasmid DNA (10 ug) and spermine (1.5 ug) were mixed in 15ul of 0.1M HEPES, pH 8.0. After 5 min of incubation DSP (25 ug in 1 mlof DMF) was added with intensive mixing. After 1 hr incubation at roomtemperature DNA was analyzed on agarose gel. In case of “DNA after”experiment, DNA (10 ug) was added after quenching DSP reaction with 0.1M glycine for 0.5 hr. Electrophoretic pattern was found similar to theone with AEPD/DSP in Example 1.

Example 5

A novel amine was used as a monomer in conjunction with DTBP fortemplate polymerization of DNA.

Methods:

The following amine was used as a monomer:

-   1. 3,3′-(N′,N″-tert-butoxycarbonyl)-N-(7-octene)-N-methyldipropyl-    ammonium bromide (compound 7, see synthesis section).

Following cross-linkers were used as co-monomers:

1. Dimethyl-3,3′-dithiopbispropionimidate (DTBP, S-S cleavable bisimidoester, Pierce)

Optimized reaction conditions with compound 6/DTBP were as follows.Plasmid DNA (pCILacZ, 10 mL of a 3.4 mg/mL stock solution, 34 mg, 103nmol nucleotide base) and compound 6 (3 mL of a 1.29 mg/mL stocksolution, 39 mg, 108 mmol) were mixed with 85 ml water and 10 ml ofbuffer solution (0.2 M HEPES, 10 mM EDTA, pH 8.0). DTBP (1.1 mL of a 100mM solution in DMF, 33.7 mg, 109 nmol) was added. After mixing, thereaction was left for 1 h in the dark at room temperature.

The pCILacZ plasmid was similarly constructed by placing the restrictiondigestion fragment of the E. coli β-galactosidase gene into the pCIvector.

Results

Agarose gel electrophoresis of the final complexes demonstratedcharacteristic retardation of the plasmid DNA. The control sample (DNAadded after reaction) did not show any retardation.

Example 6

A peptide was used as a monomer for polymerization in the presence ofDNA and this process enable the formation of complexes that expressedluciferase after transfection into cells in culture.

Methods:

NLS peptide corresponds to the 12 amino acid nuclear localizing sequenceof SV40 T antigen. This peptide was synthesized by GenosysBiotechnologies Inc with a Cysteine on each end for cross-linkingpurposes (MW=1481) Histone H1 was obtained from Sigma. The cross-linkersDSP (dithiobis[succinimidylpropionate]) and DPDPB(1,4-Di-[3′-(2′-pyridyldthio)-propionamido)]butane) were purchased fromPierce.

The NLS peptide was mixed with plasmid DNA (pCILuc) at various ratios(0.4, 0.8, 1.2, 1.6) in 20 mM HEPES pH 7.5, 1 mM EDTA at a concentrationof plasmid DNA of 0.3 mg/ml. The disulfide cleavable cross-linker DPDPBwas added to final concentrations of 2 and 6 mM and the mixtures wereincubated for 30 minutes at room temperature in the dark. Reactionproducts were analyzed by agarose gel electrophoresis and DNA wasvisualized by ethidium bromide staining. Extent of polymerization ofcationic monomers (NLS peptides) was determined on SDS-PAGE. Briefly,NLS peptide/pDNA complexes (with and without DPDPB cross-linker) wereincubated with 2.5 units DNase I for 1 hour at 37° C. to remove the DNAfrom the complexes. Following digestion, remaining protein componentswere run on a 15% SDS-PAGE and stained with coomassie blue.

All transfections were performed in 35 mm wells using 2 ug pDNA perwell. NLS peptide/pDNA complexes (with and without DPDPB cross-linker)were diluted in Opti-MEM (Life Technologies) and a fusogenic cationicpolyamine (ODAP, Mirus Corp, Madison, Wis.) was added to enhancetransfection. It is believed that this polyamine helps facilitateintracellular endosomal escape of the complexes into the cytoplasm.Pre-formed complexes were incubated with phosphate buffered salinewashed NIH3T3 cells for 4 hours at 37° C. Transfection complexes werethen removed and replaced with complete growth medium. Cells were grownfor 40–48 hours and harvested and assayed for reporter gene expression(luciferase).

For determination of luciferase activity, cells were lysed by theaddition of 100 ul for 25 mm-in-diameter plates and 200 ul for 35mm-in-diameter plates of lysis buffer (0.1% Triton X-100, 0.1MK-phosphate, 1 mM DTT pH 7.8). 20 ul of the cellular extract wereanalyzed for luciferase activity as previously reported (Wolff, J. A.,Malone, R. W., Williams, P., Chong, W., Acsadi, G., Jani, A. andFelgner, P. L. Direct gene transfer into mouse muscle in vivo. Science,1465–1468, 1990.). A Lumat LB 9507 (EG&G Berthold, Bad-Wildbad, Germany)luminometer was used.

Results:

The stepwise cross-linking of NLS peptides along the DNA templatedrastically alters the mobility of pDNA in agarose gel electrophoresis.At low peptide to pDNA ratios (0.4:1, 0.8:1) the NLS peptide/pDNA/DPDPBcomplexes migrated as a characteristic smear with several prominentbands as compared to NLS peptide/pDNA complexes without cross-linkerwhich migrated similarly to pDNA alone. At higher ratios (1.2:1, 1.6:1)the net charge of the complexes becomes positive and precipitationoccurs with or without the DPDPB cross-linker. When the pDNA is addedafter the polymerization reaction (NLS peptide/DPDPB) the agarose gelmigration pattern looks nearly identical to NLS peptide/pDNA complexeswithout cross-linker indicating that polymerization did not occurwithout the template.

To determine the degree of polymerization of the NLS peptides within theNLS peptide/pDNA/DPDPB complexes, products were analyzed on 15% SDS-PAGE(without reducing agents) after DNA removal (by DNase I digestion) asillustrated in FIG. 3. Multimers of the NLS peptide were observed onlyin the reactions when cross-linker and template (pDNA) were presenttogether with the NLS peptide monomers (panel A, lanes 6 and 7). WithDTT (50 mM) treatment prior to SDS-PAGE the NLS peptides in the templatepolymerized reactions migrated at positions corresponding to monomersonce again (panel B, lanes 6 and 7) indicating that disulfide bonds werepresent in the linkages between the monomers. Lanes: M-marker proteinstandards; 1—NLS peptide alone (7 μg); 2—DNase I alone (2.5 u); 3—NLSpeptide/pDNA; 4—NLS peptide/DPDPB (2 mM)+pDNA added after thepolymerization reaction; 5—NLS peptide/DPDPB (6 mM)+pDNA added after thepolymerization reaction; 6—NLS peptide/DPDPB (2 mM)/ pDNA; 7—NLSpeptide/DPDPB (2 mM)/pDNA. Protein staining clearly shows a ladder ofincreasing size bands indicating a stepwise polymerization of NLSpeptides with dimers appearing as the fastest migrating species. Thisladder of bands was only observed in the reactions when the cross-linker(DPDPB) was present together with the pDNA and the NLS peptideindicating that polymerization proceeded in a template dependentfashion. In addition, treatment of the complexes with the reducing agentDTT in the sample buffer completely abolished the ladder indicating thatthe ladder was a result of NLS peptide cross-linking via the disulfidecontaining DPDPB.

Polyamine mediated transfections performed with NLS peptide/pDNA/DPDPBcomplexes resulted in increased level of luciferase production ascompared to transfections with NLS peptide/pDNA alone or NLSpeptide/DPDPB polymerization prior to pDNA addition (Table 1). Forexample, with the 0.8:1 ratio of peptide to DNA compare the luciferaselevels in the control sample (9.4 million, second row, NLS+2 mM DPDPB,DNA added after reaction) to the levels in the experimental sample(365.9 million, third row, NLS+2 mM DPDPB in the presence of DNA). Theprocess of template polymerization caused a ˜40-fold increase inexpression.

TABLE 1 Luciferase expression (in light units per 35 mm well). TotalLight Units per 35 mm well × 10⁶ 0.4:1 ratio of peptide to 0.8:1 ratioCondition DNA of peptide to DNA NLS + 2 mM DPDPB, 4.4 9.4 DNA addedafter reaction NLS + 2 mM DPDPB in 10.9 365.9 the presence of DNA NLS +6 mM DPDPB in 5.7 393.1 the presence of DNAFindings:

1. Peptides can be used for template polymerization in the presence ofDNA.

2. This process enables complexes to be prepared that can transfectmammalian cells efficiently.

Example 7

Chain Polymerization using 1-Vinylimidazole (VIm) as a Monomer,2,2′-Azobis(-amidinopropane)dihydrochloride (AAP) as an Initiator andPlasmid DNA as a Template

Methods:

The conditions for template polymerization of 1-vinyl imidazole (VIm) asa monomer using 2,2′-Azobis(-amidinopropane) dihydrochloride (AAP) as aninitiator were as follows. A 400 mM stock solution of VIm (TCI America0GB01, MW 94.72, density 1.04) was prepared with sterile deionizedwater. The pH was adjusted to 6 with HCl. Then the solution was degassedwith nitrogen gas. A 200 mM stock solution of AAP (Wako 11G2606, MW271.2) was also prepared with sterile deionized water and degassed withnitrogen gas. 20 mM of plasmid DNA (pBlueRSVLux, 800 ul of 6.9 mg/ml)was mixed with 20 mM of VIm and 2 mM of AAP from the stock solutionsabove. A control sample contained VIm and AAP at the same concentrationsbut plasmid DNA was omitted. Both the experimental (VIm/AAP/DNA) andcontrol (VIm/AAP but no DNA) reactions were performed in steriledeionized water. The reactions were incubated for 2 hours at 50° C. andthen the samples were analyzed by agarose gel electrophoresis followedby ethidium bromide staining. 20 mM of plasmid DNA was added to thecontrol sample prior to loading it on the gel.

The previously described, plasmid DNA pBlueRSVLux (also known aspBS.RSVLux) was used to express the firefly luciferase reporter genefrom the Rous Sarcoma Virus (RSV) LTR promoter (Danko, I., Fritz, J. D.,Jiao, S., Hogan, K., Latendresse, J. L., and Wolff, J. A. Gene TherapyPharmacological enhancement of in vivo foreign gene expression inmuscle. volume 1, pp. 114–121, 1994). The plasmid also contains the SV40intron and poly A addition signals for proper and efficient mRNAprocessing.

Results

The agarose gel electrophoresis analysis showed that the plasmid DNA inthe control sample (DNA added after the reaction) migrated with the samepattern as the original plasmid DNA. In the experimental sample (DNApresent during the reaction), the plasmid DNA was retarded with DNAbands migrating slower than the original plasmid DNA. There was also DNAstaining material in the starting wells.

Findings:

1. A polyvinyl imidazole polymer formed in the presence of DNA and thispolymer was complexed with the DNA as evident by gel electrophoresisanalysis.

2. This polymer formed by template polymerization because the polymerdid not form if the template DNA was omitted.

Example 8

Template polymerization (caging) of large polymers

Methods:

Poly-L-lysine (hydrobromide, molecular mass from 30 to 70 kDa) (PLL) andPolyallylamine (hydrochloride)(55 kDa) (PAA) were obtained from Aldrich.Histone H1(Type III-S from Calf Thymus) was obtained from Sigma.Dimethyl 3,3′-dithiobispropionimidate(DTBP) was purchased from Pierce.The polycations were dissolved in de ionized water: PLL and H1 toconcentration 10 mg/ml and PAA to 2 mg/ml. DTBP was dissolved in H2O (30mg/ml) immediately before utilization.

DNA/polycation complexes were prepared by the rapid mixing of 37 μg ofplasmid DNA with varying amounts of polycations in 750 μl of 25 mM HEPESpH 8.0, 0.5 mM EDTA. The mixtures were kept 30 min at room temperatureand various amounts of DTBP were added. The mixtures were incubated 2hours at room temperature. 2M NaCl was added to the complexes to finalconcentration 100 mM while vigorously mixing.

Ninety degree light scattering measurements were performed using aFluorescence Spectrophotometer. The wavelength setting was 600 nm forboth the incident beam and detection of scattering light. The slits forboth beams were fixed at 2 nm. The size of the resulting complex wasdetermined by light scattering on a Brookhaven ZetaPlus particle sizer.The samples were centrifuged at 12,000 g for 7 min. The amount of DNAremaining in the supernatant was determined by measurement of theabsorbency at 260 and 280 nm.

Results:

Effect of DNA/PLL Ratio and NaCl on the Light Scattering.

PLL was added to plasmid DNA in 0.75 ml of 25 mM HEPES pH 8.0 whilevigorously mixing. The kinetics of light scattering was determinedimmediately after mixing. The turbidity of DNA/PLL complexes was wellabove that of free DNA in all range of PLL concentration As shown inFIG. 4 complex aggregation increased when molar charge ratio PLL/DNA toapproximate to land was maximal at ratio 1.17. Further increases in PLLconcentration resulted in decreasing of complex turbidity. The lightscattering did not change with time for at least for 30 min.

At low positive to negative charge ratio water-solublenonstochiometrical complexes are formed. At ratio 1 the complexes becomeinsoluble. Increasing the content of polycation may lead to the complexchanging its sign and becoming soluble again. Presumably the particlesare stabilized in solution by the positively charged loops and danglingtails of the polycation bind to the chain DNA. With increasing saltconcentration to 100 mM the charge stabilized complexes (ratio +/− morethen 1) started to aggregate (FIG. 4). The velocity of aggregationdecreased with increasing PLL/DNA ratio, but final turbidity level wasthe same for all samples.

Effect of DTBP on DNA/PLL Complexes Light Scattering.

The incubation of DNA/PLL complexes with 0.97 umol of DTBP for 2 h atroom temperature resulted in a shift of turbidity maximum to a PLL/DNAratio of 0.88 (FIG. 4). That can be explained as increasing of PLLcharge as a result of modification. Apparently, DTBP did not crosslinkPLL/DNA complexes with each other at ratio more then 1. The addition ofNaCl to a concentration of 100 mM did not change light scatteringthroughout the range of PLL concentration (FIG. 4). These resultsindicate that the addition of DTBP prevented the PLL/DNA complexes fromaggregating in 100 mM salt.

The ability to centrifuge the DNA was used as another indication ofaggregation (Table 2). All samples were centrifuged 7 min at 12,000 rpmand the amount of DNA in supernatant was determined. As showncrosslinked PLL/DNA complexes with molar ratio 4.1 and 5.9 did notprecipitate. Therefore the size of complexes were very small. Incontrast, DNA in noncrosslinked complexes were completely precipitated.

TABLE 2 The effect of DTBP on the precipitation of plasmid DNA/PPLcomplexes in the presence of 100 mM NaCl. % DNA in solution aftercentrifugation PLL/DNA ratio −DTBP +DTBP 0.585 67 77 0.879 0 0 1.171 0 02.342 0 17 4.098 0 97 5.854 0 97Effect of PLL/DNA Ratio on the Size of Complexes.

TABLE 3 The effect of varying the DNA/PLL charge (monomer:monomer) ratioon the sizes of PLL/DNA complexes with the addition of 0.97 umol DTBP.The sizes were determined by quasi elastic light scattering and numbersindicate the percent of particles <100 nm or >100 nm. Number inparentheses indicate the size (diameter in nm) of the most abundantspecies within that size range. Percentage of Particles Less or GreaterThan 100 nm no NaCl +100 mM NaCl DNA/PLL <100 nm >100 nm <100 nm >100 nm0.43  72(50)  28(200) 36(28)  64(280) 0.65  68(42)  32(196) 36(63) 64(304) 0.88 — 100(10000) — 100(10000) 1.31 — 100(10000) — 100(10000)1.74  8(65)  92(150, 680)  7(84)  93(1000) 2.61  69(33)  31(118) 11(91) 89(836) 4.12  96(43.4)  4(6580) — 100(204, 1152) 6.18 100(22.4) — —100(222, 1052) 0.43 + DTBP  29(55)  71(331) 43(31)  57(131, 374) 0.65 +DTBP  43(31)  69(339) 16(54)  84(350) 0.88 + DTBP  13(72)  87(431, 1640)21(41)  79(707, 4690) 1.31 + DTBP  87(45, 100)  3(260) — 100(10000)1.74 + DTBP  87(45, 99)  3(256) 73(55)  27(191) 2.61 + DTBP 100(32, 98)— 77(51)  23(130) 4.12 + DTBP  99(27.9)  1(6468) 69(67.6)  31(142, 2000)6.18 + DTBP  94(35.2)  6(6580) 96(68)  4(6813) 4.12 + DTBP + DTT —100(362, 8800) 6.18 + DTBP + DTT — 100(381, 8755)

In Table 3, it is clear, that PLL/DNA complexes with ratio higher then1.3 became substantially less prone to aggregate in the presence of 100mM NaCl after DTBP modification. The PLL/DNA complex stabilized reactionis intra complex crosslinking because the treatment of the modifiedPLL/DNA complexes with ratio 4.12 and 6.18 by 50 mM DTT for 1 h resultedin aggregation. In this condition the crosslinks should be cleaved butthe level of lysine modification is not changed.

Effect of PLL/DTBP Ratio on the Size and Stability of PLL/DNA Complexes.

The PLL/DNA complex in ratio 4.12 was treated by differentconcentrations of DTBP during 2 h. The size of particles without and inpresence of 100 mM NaCl was determined by quasi elastic lightscattering.

TABLE 4 The effect of varying the DTBP/PLL ratio (molar ratio of DTBP tolysine residue) on the sizes of PLL/DNA complexes. The sizes weredetermined by quasi elastic light scattering and numbers indicate thepercent of particles <100 nm or >100 nm. Number in parentheses indicatethe size (diameter in nm) of the most abundant species within that sizerange. Percentage of Particles Less or Greater Than 100 nm DTBP/PLL noNaCl +100 nM NaCl for 1 h Ratio <100 nm >100 nm <100 nm >100 nm 0 75(88) 25(586) — 100(7524) 1.01  93(44)  7(6874)  37(92)  63(600) 2.03 95(35)  5(550)  75(66)  25(190,4658) 3.05 100(52) — 100(86) —Table 4 shows that an excess of DTBP was needed for complex protectionfrom salt dependent aggregation. It should be noted that DTBP up toratio of 3.05 did not induce crosslinking between DNA/PLL particles. Forsamples with DTBP/PLL ratio 2.03 and 3.05 zeta potential were 16.16±3.23mV and 20.33±3.3 mV respectively in 25 mM HEPES pH 8.0, 100 mM NaCl.Stability of DNA/PLL Complexes to Disruption by Polyanion DextranSulfate (DS).

DNA/PLL complexes (molar ratio of 0.87, 1.74, 3.04 or 4.35 as indicatedin FIG. 5) were prepared as before but in 1 ml of buffer. 0.97 mmol ofDTBP were added. The mixtures were incubated 2 hours at roomtemperature. 10 ul of ethidum bromide (EB) (0.1 mg/ml) were added inevery sample and the samples were incubated 30 min. The aliquot portionsof DS were then added sequentially, with mixing. After each addition,the fluorescence was allowed to stabilize 30 seconds.

Addition of PLL to DNA in solution gave rapid falls in fluorescence,corresponding to complex formation. Addition of DS to pre-formedcomplexes can restore EB fluorescence and can be taken as indicator ofcomplex stability (FIG. 5). Without DTBP, the EB fluorescence rose withthe addition of DS in every ratio of PLL/DNA (FIG. 5). With DTBP, theincrease was attenuated and there was a clear influence of DTBPmodification on complex stability: the fraction of complexes could notbe disrupted in any DS concentration. The part of complexes which arestable to disruption by DS depended on PLL/DNA ratio.

DNA/PAA Complexes.

Polyallylamine (PAA) is similar to PLL and contains primary aminogroups. But average pK of PAA is low then PLL because stronger influenceof one group to another.

TABLE 5 The effect of varying the DNA/PAA ratio on the sizes of PAA/DNAcomplexes with or without the addition of DTBP. The sizes weredetermined by quasi elastic light scattering and numbers indicate thepercent of particles <100 nm or >100 nm. Number in parentheses indicatethe size (diameter in nm) of the most abundant species within that sizerange. Percentage of Particles Less or Greater Than 100 nm PAA/ no NaCl+NaCl DNA Ratio <100 nm >100 nm <100 nm >100 nm 2.17 + DTBP  7(106)77(455),16(4436) — 100(2064) 4.34 + DTBP 93(66)  7(6900) 70(94)  30(916)6.51 + DTBP 93(53)  6(163) 81(92)  19(870) 8.68 + DTBP 97(55)  3(5607)81(62)  19(182) 4.34 55(71) 45(352)  4(79)  96(863)The results in Table 5 are very similar to the results with PLL/DNAcomplexes, but large excess of polycations are required for thepreparation of stable small particles.DNA/Histone H1 Complexes.

H1 has total positive charge 55 per molecule (Mw 21.3 kDa) and can forman inter polyelectrolyte complex with DNA. In contrast to PLL and PAA,interaction of H1 with DNA leads to considerable increase of turbidityin broad range of H1 concentration. The turbidity is not changed afteraddition of 100 mM NaCl. Treatment of H1/DNA complex with charge ratio3.42 by DTBP leads to significant decrease of turbidity from 1929 to348. Following addition of NaCl causes the turbidity to increase to 458.

The centrifugation of H1/DNA complexes in buffer with 100 mM NaCl 7 minat 12,000 rpm results in precipitation of DNA, but after DTBPmodification most part of DNA stays in solution, which indicatespresence of small particles (FIG. 6). Table 6 shows that the sizes ofthe particles formed with DTBP (Table 6B) in 100 mM NaCl were muchsmaller that the particle formed without DTBP (Table 6A).

TABLE 6 The effect of varying the H1DNA charge ratio on the sizes ofPAA/DNA complexes without (A) or with (B) the addition of DTBP. Thesizes were determined by quasi elastic light scattering and numbersindicate the percent of particles <100 nm or >100 nm. Number inparentheses indicate the size (diameter in nm) of the most abundantspecies within that size range. Charge Percentage of Particles Less orGreater Than 100 nm ratio(+/ −NaCl +NaCl −) <150 nm >150 nm <150 nm >150nm A. H1/DNA-no DTBP 1.55  7(44) 29(377)64(1376) 47(25) 36(491) 3.1 6(75) 88(500)10(6000) — 92(470)8(8000) 6.2 — 10(159)90(589) —62(350)38(1825) 9.3  9(113) 91(348) —  2(208)98(1404) B. H1/DNA + DTBP1.55 19(27)6(131) 75(886) — 84(892)16(8000) 3.1 28(37) 12(168)60(603) —28(171)72(512) 6.2 — 75(166)25(1168) 47(55) 53(222) 9.3 48(117) 52(306)56(75) 44(172)Findings:

1. DNA template polymerization of large polymers yields small particlesthat do not aggregate in physiological salt solutions.

2. The ability to prepare small particles of condensed DNA that do notaggregate in a physiologic salt solution will be an extremely usefulformulation for gene transfer and therapy.

Example 9

Step DNA template polymerization using the comonomers of AEPD and aPEGylated-AEPD (Compound 18).

Step polymerization with DNA as a template was performed using thepolyamine N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD) as in Example1 except pegylated AEPD (Compound 18, N2,N2,N3,N3-tetra(PEG-aminopropyl)-AEPD, or will be referred to as AEPD-PEG) was added to thereaction mixture along with AEPD. This example teaches how to preparenon-aggregated, water-soluble particles (diameter <100 nm) of condensedDNA via the process of template polymerization.

Methods:

The mixture of the following amines was used as comonomers:

-   1. N,N′-bis(2-aminoethyl)-1,3-propanediamine (AEPD, Aldrich,    Milwaukee, Wis.).-   2. N2,N2,N3,N3-tetra(PEG-amino propyl)-AEPD (Compound 18).

The following crosslinker was used:

-   1. Dimethyl-3,3′-dithiobispropionimidate (DTBP, S-S cleavable    bisimido ester, Pierce Chemical Co.).

Optimized reaction conditions with AEPD/AEPD-PEG mixture were asfollows. Plasmid DNA (pCIluc, 10 ug), AEPD (58 ug), AEPD-PEG (5 mg) andDTBP (187 ug) were mixed in 0.5 ml of buffer solution (20 mM HEPES, 1 mMEGTA, pH 8.5). Molar ratio of total AEPD (AEPD+AEPD-PEG) to DNA base was20:1. Reaction was allowed to proceed for three hours at roomtemperature. At 10, 30, 60 and 180 min time points particle sizing wasperformed using photon correlation spectrometer as described inExample 1. The data obtained was compared with the experiment where onlyAEPD was used as monomer. Three independent samples of each templatepolymerization reaction mixture were measured and the data was expressedas mean +/− SD. Transmission electron microscopy of the AEPD/AEPD-PEGmixture samples was performed as described in Example 1.

Results:

Formation of particles of condensed DNA for AEPD/AEPD-PEG templatepolymerization mixture was confirmed both by dynamic light scatteringand by electron microscopy. Unlike AEPD alone 20:1 mixture which yieldsaggregates, AEPD/AEPD-PEG mixture resulted in increased population ofnon-aggregated individual DNA particles with the size <100 nm ofcharacteristic rod morphology.

85% of the condensed DNA stays in solution after centrifugation inmicrocentrifuge for 5 min at 12000 g. At this conditions 100% of DNAcondensed with AEPD alone (1:20 ratio) was found precipitated.

The 1:20 DNA/AEPD/DTBP reaction mixture in the presence of AEPD-PEGmolecules with protected primary amino groups (precursor of AEPD-PEGmonomer, see Example ??) resulted in formation of aggregated DNA.

Findings:

1. Template polymerization can be performed in the presence of pegylatedAEPD molecules. PEG-containing monomer was included into final condensedDNA complex.

2. Addition of pegylated PEG into standard DNA/AEPD/DTBP templatepolymerization mixture resulted in formation of non-aggregated particlesof condensed DNA via the mechanism of steric stabilization.

Example 10

Chain polymerization of compound 11 on a DNA template in an organicsolvent.

Plasmid DNA (pCI luc, 10 mg) in 660 μL water was combined with 1380 μLmethanol and 660 μL chloroform containing 144 mg of Compound 11, givinga clear solution with a 7 fold excess of positive charge. The solutionwas vortexed and allowed to stand at room temperature for 20 minutes.One half of the monophase was reserved. The remaining monophase wasseparated into two layers by the addition of an additional 375 μL water.The two resulting layers were separated. The presence of DNA in thechloroform layer was confirmed by absorbence at 260 nm. The sizes of theparticles in the chloroform layer and in the reserved monophase weremeasured on a Brookhaven ZetaPlus particle sizer. The Bligh/Dyermonophase (Bligh, E. G. and Dyer, W. J. (1959) A rapid method of totallipid extraction and purification. Can. J. Biochem. Physiol., 37,911–917.) had two groups of particles in the size range of 80–128 nm and7000–10000 nm. The chloroform layer showed one group of particles with asize range of 4–7 nm, however the signal intensity was low.

The Bligh/Dyer monophase and the chloroform layer were polymerized inthe presence of 1% AIBN (Aldrich Chemical Company) for 1 hour at 55° C.Particle size was measured after the polymerization. The Bligh/Dyermonophase contained one population of particles with a size range of700–1000 nm. The chloroform layer contained one population of particleswith a size range of 340–400 run. The polymerized reaction products wereanalyzed on a SDS-PAGE gel (Novex, 10–20% tricene). Approximately 50 μgof the polymerized reactions and a control consisting of 7 μg DNA and 50μg compound 11 without polymerization were loaded onto the gel andvisualized with coumassie staining. A smear of high-molecular weightpolymer beginning in the well was observed in both of the polymerizationreactions. The control exhibited one band of low molecular weight.

Example 11

Methods:

Dextran sulfate (Mw=500 000, DS) was obtained from Sigma.Polymethacrylic acid sodium salt (pMAA, Mw=9 500) was obtained fromAldrich. Caged DNA particles were prepared as described in example“Template polymerization (caging) of large polymers” with DTBP ascross-linking agent. Zeta-potentials of the obtained particles weremeasured using Zeta-Plus Photon Correlation Spectrometer (BrookhavenInstruments Corp.). Ninety degree light scattering measurements and TOTObinding assay were performed using Fluorescence Spectrophotometer. TOTOassay was used to assess the degree of DNA condensation (Wong F M.Reimer D L. Bally M B, Cationic lipid binding to DNA: characterizationof complex formation. Biochemistry. 35(18):5756–63, 1996).

Results:

After the obtaining soluble particles of positevly-charged caged DNA/PLLcomplexes their surface was rendered negatively charged by complexing itwith the excess of polyanion. It was found that upon addition ofpolyanion solution to soluble DNA/PLL complex the net charge of thetriple complex can be changed to the opposite at the certainconcentration of the polyanion

TABLE 7 DS added Z-potential Z-potential ug uncaged caged 0 29.55 28.550 33.68 19.73 100 −17.25 −14.66 150 −22.45 −13.73 200 −19.84 −18.21 300400 500 −19.21 −14.33

Table 7. Zeta potential of caged and non-caged DNA/PLL (1:6) complexesafter addition of dextran-sulfate (DS). Complexes were prepared with 30ug of DNA and 114 ug of PLL and caged with 240 ug of DTBP for 2 hrs.

Triple complexes formed at 150 ug of DS were tested on solubility atphysiological salt. It was found that 60% of I90 stays in the solutionafter 5 min centrifugation at 12 000 g for both caged and uncagedcomplexes. Particle sizing using dynamic light scattering demonstrated80% particles <150 nm in diameter at these conditions.

TABLE 8 TOTO signal, TOTO signal DS added, % of native DNA % of nativeDNA ug uncaged caged 0 4 3 50 3 2 100 14 7 200 49 27 300 43 22 400 39 20500 38 20

Table 8. DNA condensation after complexing DNA/PLL complexes withdextran sulfate.

TOTO assay (Table. 8) demonstrated that DNA stays condensed afterformation of negatively charged triple complex though some partialdecondensation occured. Caged complex was found more resistant todecondensation during recharging (80% condensation preserved at 400–500ug DS added).

It is possible to recharge DNA/PLL complexes with other polyanions. Thesimilar data were obtained with polymethacrylic acid (not shown).

Syntheses of Compounds

Materials and Methods: ¹H-NMR spectra were recorded on a Bruker AC 250or a Bruker AC 300 spectrophotometer with chemical shifts given in partsper million downfield from an internal standard of tetramethylsilane.Diamino-N-methyldipropylamine (Aldrich Chemical Co.), Boc anhydride(Aldrich Chemical Co.), triethylamine (Aldrich Chemical Co.),trifluoroacetic anhydride (Aldrich Chemical Co.), 9-bromo-1-nonanol(Aldrich Chemical Co.), acryloyl chloride (Aldrich Chemical Co.),3-bromopropylamine hydrobromide (Aldrich Chemical Co.), 7-bromo-1-octene(Aldrich Chemical Co.), trimethylamine (25% solution in water) (AldrichChemical Co.), methyl iodide (Aldrich Chemical Co.),N,N,N′N′-tetramethyl-propane diamine (Aldrich Chemical Co.),N,N,N′,N′,N″-pentamethylethylentriamine (Aldrich Chemical Co.) were usedas supplied. All solvents were obtained from Aldrich Chemical Co. Allanhydrous solvents were obtained from Aldrich Chemical Co. inSure/Seal_containers.

3,3′-(N′,N″-tert-butoxycarbonyl)-N-methyldipropylamine (1).3,3′-Diamino-N-methyldipropylamine (0.800 mL, 0.721 g, 5.0 mmol) wasdissolved in 5.0 mL 2.2 N sodium hydroxide (11 mmol). To the solutionwas added Boc anhydride (2.50 mL, 2.38 g, 10.9 mmol) with magneticstirring. The reaction mixture was allowed to stir at room temperatureovernight (approximately 18 hours). The reaction mixture was made basicby adding additional 2.2 N NaOH until all t-butyl carboxylic acid was insolution. The solution was then extracted into chloroform (2×20 mL). Thecombined chloroform extracts were washed 2×10 mL water and dried overmagnesium sulfate. Solvent removal yielded 1.01 g (61.7%) product as awhite solid: ¹H-NMR (CDCl₃) d 5.35 (bs, 2H), 3.17 (dt, 4H), 2.37 (t,4H), 2.15 (s, 3H), 1.65 (tt, 4H), 1.45 (s, 18H).

3,3′-Trifluoroacetamidyl-N-methyldipropylamine (2).3,3′-Diamino-N-methyldipropylamine (0.504 mL, 436 mg, 3.0 mmol) andtriethylamine (0.920 mL, 6.6 mmol) were dissolved in 20 mL dry methylenechloride. The solution was chilled on an ice bath. Trifluoroaceticanhydride (0.932 mL, 1.39 g, 6.6 mmol) dissolved in 40 ml dry methylenechloride was added dropwise with magnetic stirring over a period ofapproximately 20 minutes. The reaction was allowed to come to roomtemperature and to stir overnight (approximately 18 hours). The reactionmixture was washed 2×15 mL 2% sodium bicarbonate, 2×15 mL water, anddried over magnesium sulfate. Solvent removal yielded 763 mg (67.9%)product as a yellow oil: ¹H-NMR (CDCl₃) d 8.20 (bs, 2H), 3.45 (dt, 2H),2.47 (t, 2H), 2.20 (s, 3H), 1.75 (tt, 2H).

9-Bromononacrylate (3). 9-Bromo-1-nonanol (0.939 g, 4.0 mmol) wasdissolved in 4.0 mL anhydrous diethyl ether in a flame dried 10 mL r.b.flask under dry nitrogen. Sodium carbonate (6.36 g, 6.0 mmol) was addedto the reaction mixture. Acryloyl chloride (0.356 mL, 0.397 g, 4.2 mmol)dissolved in 3.5 mL anhydrous ether was added dropwise over a period ofapproximately 10 minutes. The reaction mixture was allowed to come toroom temperature and stir for two days. The reaction mixture was dilutedto 40 mL with ether and washed 3×10 mL 2% sodium bicarbonate to removeunreacted acryloyl chloride. The organic layer was dried over magnesiumsulfate and passed through a short (approximately 7 g) alumina column toremove unreacted alcohol. Solvent removal yielded 390 mg (35.2%) productas a clear liquid: ¹H-NMR (CDCl₃) d 6.40 (dd, 1H), 6.12 (dd, 1H), 5.82(dd, 1H), 4.15 (t, 4H), 3.40 (t, 2H), 1.85 (dt, 2H), 1.65 (dt, 2H), 1.35(m, 10H).

3-Bromo-1-(trifluoroacetamidyl)propane (4). 3-Bromopropylaminehydrobromide (2.19 g, 10.0 mmol) and triethylamine (1.67 mL, 12.0 mmol)were dissolved in 60 mL dry methylene chloride. The solution was chilledon an ice bath. Trifluoroacetic anhydride (1.69 mL, 2.51 g, 12.0 mmol)dissolved in 60 mL dry methylene chloride was added dropwise overapproximately 20 minutes. The reaction was allowed to come to roomtemperature and was stirred overnight (approximately 18 hours). Thereaction mixture was washed 1×10 mL 2% sodium bicarbonate, 1×10 mLwater, and dried over magnesium sulfate. Solvent removal yielded 2.07 g(88.5%) product as a white powder: ¹H-NMR (CDCl₃) d 6.70 (bs, 1H), 3.55(dt, 2H), 3.45 (t, 2H), 2.17 (tt, 2H).

1-Octene-7-trimethylammonium bromide (5). 7-Bromo-1-octene (0.168 mL,191.2 mg, 1.00 mmol) was combined with trimethylamine (2.40 mL 25%solution in water). The mixture was incubated at 50 C_ on a rotaryshaker for 18 hours. Solvent removal and recrystalization fromacetone/diethyl ether yielded 191.6 mg (76.6%) product as white plates:¹H-NMR (CDCl₃) d 5.75 (m, 1H), 5.00 (m, 2H), 3.60 (m, 2H), 3.45 (s, 9H),2.05 (m, 2H), 1.75 (m, 2H), 1.40 (m, 6H).

3,3′-(N′,N″-tert-butoxycarbonyl)-N-(7-octene)-N-methyldipropyl-ammoniumbromide (6). Compound 1 (86.3 mg, 0.25 mmol) was combined with7-bromo-1-octene and dissolved in 0.050 mL methyl sulfoxide. Thereaction mixture was incubated at 55 C_ for 18 hours. The viscousreaction mixture was triturated with ether twice. The remaining oil wasrecrystalized from chloroform/ether to yield 55.3 mg (48.7%) product aswhite crystals: ¹H-NMR (CDCl₃) d 5.75 (m, 3H), 4.95 (m, 2H), 3.55 (m,4H), 3.30 (m, 6H), 3.15 (s, 3H), 2.05 (m, 4H), 1.97 (m, 2H), 1.70 (m,2H), 1.45 (s, 18H), 1.35 (m, 6H).

3,3′-diamine-N-(7-octene)-N-methyldipropylammonium bromide (7). Compound6 was combined with 0.350 mL ethyl acetate, 0.150 mL methanol, and 0.150mL 12 N hydrochloric acid. The reaction mixture was stirred at roomtemperature for 2.5 hours, during this time the reaction becamehomogenous. Solvent was removed and the product was precipitated from asmall amount of methanol with ether to yield 36.0 mg (95.2%) product asa colorless oil: ¹H-NMR (CD₃OD) d 5.85 (m, 1H), 5.00 (m, 2H), 3.55 (m,4H), 3.45 (m, 2H), 3.20 (s, 3H), 3.15 (t, 4H), 2.25 (m, 4H), 2.10 (m,2H) 1.85 (m, 2H), 1.50 (m, 6H).

3,3′-(N′,N″-tert-butoxycarbonyl)-N,N-dimethyldipropylammonium bromide(8). Compound 1 (75.0 mg, 0.217 mmol) was dissolved in 0.5 mL dry ether,ethyl alcohol was added drop-wise until compound 1 was completelydissolved. The reaction solution was chilled on an ice bath and purgedwith dry nitrogen. Methyl iodide (0.021 mL, 33.7 mmol) was added, andthe reaction mixture was stirred at 4 C_ for 18 hours. Poured reactionmixture into water and washed with ether. After removal of water theproduct was dissolved in chloroform, decolorized with activatedcharcoal, and dried with magnesium sulfate. Solvent removal yielded 92.0mg (87.0%) product as a yellow oil: ¹H-NMR (CDCl₃) d 5.50 (bs, 2H), 3.60(m, 4H), 3.30 (s, 6H), 3.25 (m, 4H), 2.07 (m, 4H), 1.45 (s, 18H).

3,3′-Diamino-N,N-dimethyldipropylammonium bromide (9). Compound 8 (92.0mg, 0.189 mmol) was dissolved in 0.200 mL ethyl acetate and 0.150 mL 12N hydrochloric acid. The reaction mixture was stirred at roomtemperature for 1 hour. Solvent was removed and the oily residue wastriturated three times with ether. The remaining product was dried invacuo yielding 43.9 mg (100%) product as a yellow waxy solid: ¹H-NMR(CD₃OD) d 3.55 (m, 4H), 3.20 (s, 6H), 3.20 (t, 4H), 2.22 (m, 4H).

N,N′-Dinonacrylate-N,N,N′,N′-tetramethylpropanediammonium bromide (10).N,N,N′N′-tetramethylpropane diamine (0.0252 mL, 0.15 mmol) and compound3 (131 mg, 0.148 mmol) were dissolved in 0.150 mL dimethylformamide. Thereaction mixture was incubated at 50 C_ for 5 days. The product wasprecipitated from the reaction mixture by the addition of ether. Theresulting solid was collected and recrystalized twice from ethanol/etheryielding 56.9 mg (55.4%) product as white crystals: ¹H-NMR (CDCl₃) d6.40 (dd, 2H), 6.15 (dd, 2H), 5.85 (dd, 1H), 4.15 (t, 4H), 3.88 (m, 4H),3.52 (m, 4H), 3.40 (s, 12H), 2.75 (m, 2H), 1.82 (m, 4H), 1.65 (m, 4H),1.35 (m, 20H).

N,N′,N″-Trinonacrylate-N,N,N′,N′,N″-pentamethyldiethylentriammoniumbromide (11). N,N,N′,N′,N″-pentamethylethylentriamine (0.031 mL, 0.15mmol) and compound 3 (187 mg, 0.675 mmol) were dissolved in 0.150 mLdimethylformamide. The reaction mixture was incubated at 50 C_ for 5days. The product was precipitated from the reaction mixture by theaddition of ether. The resulting solid was collected and recrystalizedfrom ethanol/ether yielding 36.6 mg (24.3%) product as white crystals:¹H-NMR (CDCl₃) d 6.40 (dd, 3H), 6.15 (dd, 3H), 5.83 (dd, 3H), 4.15 (t,6H), 3.95 (m, 4H), 3.60 (m, 4H), 3.40 (s, 15H), 3.17 (m, 6H), 1.70 (m,12H), 1.35 (m, 30H).

3,3′-Trifluoroacetamidyl-N-nonacrylate-N-methyldipropylammonium bromide(12). Compound 2 (233 mg, 0.691 mmol and compound 3 (282 mg, 1.01 mmol)were dissolved in 0.200 mL dimethylformamide. The reaction mixture wasincubated at 50 C_ for 4 days. The product was precipitated from thereaction mixture by the addition of ether. The resulting oil wastriturated 3× with ether. The oil was dried in vacuo yielding 385.5 mg(90.8%) product as a clear oil: ¹H-NMR (CDCl₃) d 9.05 (bs, 2H), 6.35(dd, 1H), 4.15 (t, 2H), 3.50 (m, 8H), 3.20 (m, 2H), 3.15 (s, 3H), 2.20(m, 4H), 1.70 (m, 4H), 1.30 (m, 10H).

3,3′-(N′,N″-tert-butoxycarbonyl)-N-(3′-trifluoroacetamidylpropane)-N-methyldipropylammoniumbromide (13). Compound 1 (100.6 mg, 0.291 mmol) and compound 4 (76.8 mg,0.328 mmol) were dissolved in 0.150 mL dimethylformamide. The reactionmixture was incubated at 50 C_ for 3 days. TLC (reverse phase;acetonitrile: 50 mM ammonium acetate pH 4.0; 3: 1) showed 1 major and 2minor spots none of which corresponded to starting material.Recrystalization attempts were unsuccessful so product was precipitatedfrom ethanol with ether yielding 165.5 mg (98.2%) product and minorimpurities as a clear oil: ¹H-NMR (CDCl₃) d 9.12 (bs, 1H), 5.65 (bs,2H), 3.50 (m, 8H), 3.20 (m, 4H), 3.15 (s, 3H), 2.20 (m, 2H), 2.00 (m,4H), 1.45 (s, 18H).

3,3′-(N′,N″-tert-butoxycarbonyl)-N-(3″-aminopropane)-N-methyl-dipropylammoniumbromide (14). Compound 13 (1.09 g, 1.88 mmol) was dissolved in 10 mLmethanol and 1.0 mL water. Sodium carbonate (1.00 g, 9.47 mmol) wasadded, and the reaction mixture was stirred at room temperature for 18hours. Sodium carbonate and solvent were removed leaving a clear oilwhich was triturated 3× with ether acuum drying yielded 898.2 mg (98.8%)product as a white solid. TLC (reverse phase; acetonitrile: 50 mMammonium acetate pH 4.0; 1:3) gave 1 spot rf=0.54. ¹H-NMR (D₂O) d 3.55(m, 6H), 3.27 (m, 4H), 3.05 (s, 3H), 2.87 (m, 2H), 1.97 (m, 6H), 1.45(s, 18H).

N₁,N₄-(tert-butoxycarbonyl)-bis(2-aminoethyl)-1,3-propanediamine (15)AEPD (275 mg, 1.72 mmol) was dissolved in 5.0 mL tetrahydrofuran andchilled to 0° C. on an ice bath. BOC-ON (800 mg, 3.25 mmol, AldrichChemical Co.) dissolved in 3 mL tetrahydrofuran was added dropwise withmagnetic stirring over approximately 15 minutes. The ice bath wasremoved and the stirring reaction mixture was allowed to come to roomtemperature. After 2 hours the solvent was removed on a rotaryevaporator, and the residue was dissolved in 20 mL chloroform. Thechloroform was washed with 2 N sodium hydroxide. The chloroform layerwas then extracted with 0.1 N hydrochloric acid. The acid layer was thenmade basic by the addition of 2 N sodium hydroxide and the product wasback extracted into chloroform. The chloroform was dried over magnesiumsulfate. Solvent removal afforded 288 mg product (49.2%). ¹H-NMR (CD₃OD)∂ 3.15 (t, 4H), 2.65 (m, 8H), 1.70 (m, 2H), 1.45 (s, 18H).

N₂,N₂,N₃,N₃-(3′-trifluoroacetamidylpropane)-N₁,N₄-(tert-butoxycarbonyl)-bis(2-aminoethyl)-1,3-propanediammoniumdibromide (16) Compound 15 (33.0 mg, 91.7 μmol) and3-Bromo-1-(trifluoroacetamidyl)propane (128 mg, 547 μmol) were combinedin 200 μL dimethylformamide and incubated at 55° C. for 24 hours. TLC(silica: 90% acetonitrile, 10% 50 mM ammonium acetate pH 4.0) showed amixture of products. Additional 3-Bromo-1-(trifluoroacetamidyl)propane(100 mg, 427 μmol) in 200 μL dimethylformamide and incubated anadditional 24 hours. TLC showed 2 spots at rf of 0.51 and 0.58 in theabove system when developed with dragendorffs reagent (Sigma ChemicalCo.) Precipitation with diethyl ether yielded 56.0 mg product (53%). Thefinal product may be a mixture of tri-alkylated AEPD and tetra-alkylatedAEPD. Full characterization and purification is in progress.

N₂,N₂,N₃,N₃-(3′-aminopropane)-N₁,N₄-(tert-butoxycarbonyl)-bis(2-aminoethyl)-1,3-propanediammoniumdi-trifluoroacetate (17) Compound 16 (28.0 mg, 24.7 μmol) dissolved in 1mL 6:4 methanol water along with calcium carbonate (104 mg, 1.0 mmol).The reaction mixture was stirred at 60° C. for 3 hours. TLC (silica: 90%acetonitrile, 10% 50 mM ammonium acetate pH 4.0) indicated completion ofreaction with all material remaining at the origin. Product was isolatedafter removal of calcium carbonate by filtration to yield 13.0 mg(75.1%).

N₂,N₂,N₃,N₃-(3′-PEG₅₀₀₀aminopropane)-N₁,N₄-bis(2-aminoethyl)-1,3-propanediammoniumdi-trifluoroacetate (18) Compound 17 (13 mg, 18.6 μmol) and0-[2-(N-succinimidyloxycarbonyl)-ethyl]-O′-methylpolyethylene glycol5,000 [NHS-Peg] (180 mg, 36 μmol) in 0.5 mL dimethylformamide. Thereaction was stirred for 30 minutes. The reaction mixture was checkedfor the presence of primary amines by spotting on TLC plate anddeveloping with ninhydrin spray. Primary amines were still present soadditional NHS-Peg (80 mg, 16 μmol) was added. The reaction mixture wasagain screened for the presence of primary amines, none were found to bepresent. The reaction was stopped by precipitation with diethyl ether.The precipitate was washed 2× with diethyl ether, and dried under vacuumto yield 198 mg product. The product was dissolved in 2 mLtrifluoroacetic acid, and incubated 20 minutes to remove the BOCprotecting groups. The trifluoroacetic acid was removed under a streamof nitrogen. The residue was dried under vacuum to yield 198 mg productas an off-white solid. The presence of free amino groups after theremoval of the BOC protecting groups was confirmed by a positiveninhydrin test. The final product should contain approximately 3 Pegchains per molecule as determined by the amount of NHS-Peg used inreaction.

The foregoing is considered as illustrative only of the principles ofthe invention. Furthermore, since numerous modifications and changeswill readily occur to those skilled in the art, it is not desired tolimit the invention to the exact construction and operation shown anddescribed. Accordingly, all suitable modifications and equivalents fallwithin the scope of the invention.

1. A method for delivering a polyion to a cell, comprising: a)noncovalently associating polymers with the polyion in a solutionoutside a cell; b) forming covalent linkages between moieties of thepolymers in the presence of the polyion without chemically modifying thepolyion, thereby forming a complex; and, c) delivering the complex tothe cell.
 2. The method of claim 1 wherein the polymers are selectedfrom the group consisting of polycations and polyanions.
 3. The methodof claim 1 further comprising attaching molecules to the polymers. 4.The method of claim 3 wherein the molecules consist of targeting groups.5. The method of claim 4 wherein the targeting groups are selected fromthe group consisting of nuclear localizing signals, ligands that bind tocellular receptors, and releasing signals.
 6. The method of claim 3wherein the molecules are selected from the group consisting ofamphipathic, hydrophobic and hydrophilic compounds.
 7. The method ofclaim 1 further comprising: mixing the complex with a second polymer ofopposite charge to the polymers of claim
 1. 8. The method of claim 7wherein the net charge of the complex is changed.
 9. A method of makinga complex for delivery to a cell, comprising: a) non-covalentlyassociating a molecule with a polyion in a solution outside a cell; and,b) covalennly modifying the molecule in the presence of the polyionwithout chemically modifying the polyion thereby providing a deliverablecomplex.
 10. The method of claim 1 wherein the complex is in ahypertonic solution.
 11. A method of making a stabilized complex fordelivery to a cell, comprising: a) non-covalently associating polymerswith a polyion outside a cell to form a complex; b) forming covalentlinkages between moieties of the polymers in the presence of the polyionwithout chemically modifying the polyion, thereby forming a stabilizedcomplex.